The authors thank members of the Lwigale laboratory for critique of the manuscript. SLG is supported by a Ruth L. Kirschstein NRSA Fellowship from the National Eye Institute (F32 EY02167301). PYL is supported by the National Eye Institute (EY018050).

1. Incubate chick and quail eggs to the desired stage
For HH9 embryos, typical incubation times range from 29-33 hours at 38 &deg;C.63
<ol>
	<li>Wash any debris from the eggs with tepid water.</li>
	<li>Arrange chicken eggs on tray horizontally. Mark the side facing up with pencil; this will correspond to the region where the embryo will be localized. Incubate quail eggs blunt end up.</li>
	<li>Place in 38 &deg;C humidified incubator. Turn on rocking function.</li>
</ol>
2. Prepare eggs for windowing and dissection
<ol>
	<li>Remove eggs from incubator, and sterilize the tops with 70% ethanol. It is best to spray on the ethanol and wipe it off quickly with a paper towel or Kimwipe to avoid any absorption of ethanol through the eggshell.</li>
	<li>Place chicken egg onto an individual egg holder (we use a Petri dish lined with folded paper towels, e.g., "Kimwipes"). Using AA forceps, tap a small hole into the upper surface of the eggshell at the pointed end of the egg.</li>
	<li>Remove 1.5-3mL of light albumin from the chicken egg with an 18&frac12; G hypodermic needle and 5 mL syringe. It is best to insert the needle vertically into the hole, with the bevel facing the pointy end of the egg (Figure 2A). This way, when albumin is withdrawn, there is little risk of accidentally damaging the yolk via suction. Discard the albumin. This step will lower the yolk and embryo within the egg to allow for a window to be opened in the eggshell.</li>
	<li>Wipe the hole with 70% ethanol as described above and seal it with Scotch tape. It is important that the eggshell surrounding the hole is completely devoid of albumin and dry or the adhesive will not seal.</li>
	<li>With the AA forceps, tap another hole into the marked upper surface of the horizontal egg, not quite at the apex. Take care not to let the forceps go too far through the eggshell to avoid damaging the yolk or embryo.</li>
	<li>Insert one side of the curved iris forceps into the hole parallel to the eggs shell. Pinching down on the shell, working in a circular motion to break a ~2 cm in diameter window in the chicken egg shell. Discard the removed eggshell. Alternatively, cover the upper surface of the egg with packing tape and use a pair of scissors to cut out the window. This minimizes the chance of eggshell debris falling into the egg. The embryo should appear as an opaque disc on top of the yolk. Discard any unfertilized eggs (identified by small white spot on surface of yolk, or absence of blastoderm).</li>
	<li>Inject a small amount of India ink (diluted 1:10 in sterile Ringer's solution containing "Pen/Strep" antibiotic: final concentration 100 &mu;g/mL penicillin and 100 &mu;g/mL streptomycin) beneath the blastoderm to assist with staging the embryo. Use a 1 mL syringe and 26&frac12; G hypodermic needle, bent into a 45&deg; angle at the base of the needle, with the bevel facing up; alternatively, one may use a pulled glass Pasteur pipet and mouth pipetting apparatus (wear eye protection when bending hypodermic needle or breaking glass needle). Puncture the yolk membrane outside the perimeter of the blastoderm and slide the tip of the needle beneath the embryo, close to the surface of the yolk but not in the embryonic layers (Figure 2B). Inject just enough ink to outline the embryo, then carefully withdraw the needle (Figure 2C). Injecting too much ink can lead to embryo death. It is important not to inject any air bubbles beneath the embryo, as this can be a source for contamination. NB: All Ringer's solution used in this procedure is sterilized and contains Pen/Strep as defined above. Sterile phosphate buffered saline (PBS) is also an acceptable alternative to Ringer's solution in all steps of this protocol.</li>
	<li>Stage the embryo according to Hamburger and Hamilton63 and record the stage on the eggshell in permanent ink or pencil. Stages are best assessed under stereomicroscopy, with fiber optic "gooseneck" light sources which have a limited heat load.</li>
	<li>Apply 2-3 drops warm sterile Ringer's solution to the surface of the embryo to prevent dehydration and contamination. Temporarily seal the window with parafilm stretched over the surface of the egg.</li>
	<li>Repeat steps 2.2-2.9 with all chick eggs before beginning the transplant experiments.</li>
	<li>Since quail embryos are only used for donor tissues, in ovo steps 2.2-2.9 may be omitted. For ex ovo preparation of donor embryos, quail eggs are incubated blunt side up and opened in this region with curved forceps to reveal the embryo. Using dissecting scissors, make four cuts in the shape of a square through the vitelline membrane and blastoderm outside the embryo (make sure that all cuts meet). Using a curved iris forceps, gently grasp one cut edge and lift the embryo from the egg and transfer it to Ringers solution in a Petri dish. Alternatively, one may use the curved iris forceps to lift the excised embryo from beneath, or an embryo spoon or plastic transfer pipette cut through the wider part of the cylinder, may be used to transfer the embryo out of the egg. Gently remove the vitelline membrane with #5 forceps. Collect the remaining quail embryos as above, and stage them under a dissecting microscope.</li>
</ol>
3. Prepare the host embryo to receive the graft
<ol>
	<li>Remove parafilm from the host (chick) embryo and using a sharpened tungsten needle or #5 forceps, tear a small hole in the vitelline membrane at the desired region of the neural tube (Figure 3A).</li>
	<li>Add a drop of Ringer's solution to the embryo. Take care to keep the embryo completely submerged in Ringer's solution during the entire procedure to prevent dehydration of the tissue. Keep adding drops of Ringer's solution to the surface of the embryo via transfer pipette if necessary.</li>
	<li>Using a pulled glass needle, carefully make rostral and caudal transverse incisions corresponding to the length and region of interest in the dorsal neural tube. (Pulled glass needles are generated from silicon glass capillary tubes which have been pulled under heat with a needle pulling apparatus.) For unilateral grafts the incisions should only extend to the lumen of the dorsal neural tube, and for bilateral grafts, make the incisions across the entire dorsal neural tube. Then cut bilaterally between the dorsal neural tube and the paraxial mesoderm.</li>
	<li>Carefully separate the excised explant from the neural tube then remove it from the embryo by aspirating into a glass micropipette (Figure 3B).</li>
</ol>
4. Prepare the donor graft tissue
<ol>
	<li>Choose a stage-matched quail embryo. Hold the embryo down with AA forceps, remove the vitelline membrane, and excise a similar-sized region of neural tube as described above (3.3-3.4) (Figure 3A'-B'). Depending on the needs of your experiment, this region may be a complementary region of the host (chick) neural tube or from a different region along the rostro-caudal axis. (NB: for larger regional grafts, including complete neural tube grafts, researchers may wish to add a protease digestion at this step to remove any adherent mesenchymal tissue from the donor tissue prior to transplantation. However, for small dorsal neural tube grafts at early stages, as is detailed here, protease digestion is not necessary as the neural tube is easily separated from adjacent mesenchyme by strictly surgical techniques.)</li>
</ol>
5. Graft the tissue
<ol>
	<li>Aspirate the donor explant into a glass micropipette containing Ringer's solution. Be careful not to introduce any bubbles into the pipette.</li>
	<li>Transfer the explant adjacent to the excised region of the chick host. Using pulled glass needle, orient the explant and gently guide it into the ablated region (Figure 3C).</li>
	<li>Carefully add a few drops of Ringer's solution to the embryo to prevent dehydration. Take care to avoid dropping liquid directly on the graft site as this may dislodge the graft.</li>
	<li>Seal the window with packing tape. Make sure that the tape seals the entire region of the windowed egg. This prevents dehydration and contamination of the embryo during subsequent incubation.</li>
	<li>Return the egg to the incubator until the desired stage. Make sure that the "rocking" function is turned off while incubating the chimeras. Eggs may be gently turned by hand two times a day as this may increase their viability if later stages of development are targeted.</li>
</ol>
6. Prepare chimeric embryos for sectioning
<ol>
	<li>At the experimental endpoint, cut away the tape covering the window using fine scissors.</li>
	<li>Grasp the edge of the blastoderm with curved iris forceps (easiest to do with serrated tips), then cut the embryo away from the yolk with 4 large cuts outside the boundaries of the blastoderm.</li>
	<li>Transfer the embryo to a Petri dish containing Ringer's solution. Take care to minimize exposure of the embryo to air in order to avoid dehydration of the tissue.</li>
	<li>Under stereomicroscopy, gently separate the vitelline membrane from the blastoderm using #5 forceps.</li>
	<li>Carefully arrange the embryo in the dish so that the embryo is in the same orientation as it was in the egg, with surrounding membranes laid out flat.</li>
	<li>Use dissecting needles made from tungsten wires64 to cleanly remove the surrounding blastoderm from the embryo by arranging the needle at the boundary of the embryo and extraembryonic tissues, with the length of the tungsten wire parallel to the bottom of the dish. Use a gentle sawing motion to cut through the tissues.</li>
	<li>Carefully remove the amnion with #5 forceps. At this point you may, wish to either further dissect the region of your interest, or leave the embryo whole.</li>
	<li>Transfer the embryo to ice cold 4% paraformaldehyde and rock overnight at 4 &deg;C.</li>
	<li>Prepare samples and embed for cryo- or paraffin sectioning.</li>
</ol>
Trace the grafted tissue within the host embryo. There are several techniques for identifying quail tissue within a chick embryo, including detection of quail nucleoli by hematoxylin staining (quail nuclei have very large, darker staining inclusions than chick nuclei), the Feulgen-Rossenbeck reaction, acridine-orange or biz-benzamide stain combined with electron microscopy, or immunolabeling for quail-specific antigens3,47,65,66. Here we use QCPN antigen and standard whole-mount or section immunofluorescence techniques to identify quail neural crest cells in quail-chick chimeras (Figure 4). This technique provides the most flexibility in experimental design, as the QCPN immunofluorescence may also be combined with other antibodies to identify differentiated cells derived from the donor (quail) tissue. Use standard whole-mount or section immunofluorescence techniques to label quail-derived cells in chimeras.
7. Representative Results
A representative image of the grafted region of the neural tube after 6h of re-incubation (to HH11) shows expected incorporation of the grafted donor (quail) tissue into the host (chick) neural tube (Figure 4A). Embryos showing incomplete integration of the graft, or asymmetric development of the cranial region or somites after reincubation should be discarded.
Cross section through the grafted region at HH11 show NCCs labeled with HNK-1 migrating laterally away from the neural tube. Quail cells contributing to the NCC migratory stream, and to the neural tube, are clearly labeled with QCPN (Figure 4B).
At later stages, quail NCC-derived cells can be traced to their final target tissue. QCPN labeled cells are interspersed within the chick embryo mesenchyme of the maxillary process at E5 (Figure 4C).
The QCPN antibody can be easily combined with other antibodies to examine the differentiation of quail-derived NCCs in the host environment. Quail NCC-derived trigeminal sensory neurons are labeled by QCPN and Tuj1 antibodies (Figure 4D).
<img alt="Figure 1" src="/files/ftp_upload/3622/3622fig1.jpg" /> 
Figure 1. Overview of experimental procedure and necessary instruments. A) Host (chick) and donor (quail) embryos should be stage matched. To label the NCC stream that contributes to the trigeminal ganglion and maxillo-mandibular region, HH9 is ideal. At HH9, the neural tube is beginning to close in the rostral region, but is not yet completely sealed. The gray line in the diagrams represents the midline of the closing neural tube. The dotted white lines indicate the cut lines for excising a midbrain region of the right side of the neural tube from host and donor embryos. The excised region of the host embryo is discarded and the donor tissue transplanted in to generate the chimeric embryo. B) Necessary instruments include: a) 5mL syringe with 18&frac12; G hypodermic needle, b) 1 mL syringe with 26&frac12; G hypodermic needle bent at 45&deg; angle, c) India ink, d) clear packing tape, e) glass pipette with mouth pipetting apparatus, f) 60 mm-Petri dish, g) AA forceps, h) curved iris forceps with serrated tips, i) #5 forceps, j) fine scissors, k) sharpened tungsten wire, l) pulled glass needle, m) parafilm squares.
<img alt="Figure 2" src="/files/ftp_upload/3622/3622fig2.jpg" /> 
Figure 2. Preparation of eggs and embryos. A) Cross-sectional diagram of the egg, including ideal insertion point of 18&frac12; G hypodermic needle for withdrawal of light albumin. B) After withdrawing ~3mL of light albumin, the yolk and embryo lowers within the egg, allowing the researchers to cut a "window" (arrows) in the eggshell to access the embryo. India ink, diluted 1:10 in sterile Ringer's solution, can then be injected beneath the blastoderm to provide contrast for easy staging of the embryos. C) Chick embryo after inking.
<img alt="Figure 3" src="/files/ftp_upload/3622/3622fig3.jpg" /> 
Figure 3. Schematic and examples of HH stage 9 donor and host embryos at each step of the grafting procedure. A) Chick host embryo. Dashed lines in diagram indicate where the vitelline membrane should be torn to access the cranial region for grafting. Once torn the triangular flap of vitelline membrane should be peeled caudally. Box in image indicates inset used in (B-C). A') Quail donor embryo. Image is of donor embryo excised from egg and placed into Petri dish for dissection of the graft tissue. Box in image indicates inset used in (B'). Dashed lines in schematic diagram (A') indicate where the vitelline and yolk membranes should be cut in order to remove the embryo from the egg. B) Host embryo with unilateral midbrain region of neural tube excised (white arrows) awaiting graft. Dotted line in schematic diagram indicates where cuts should be made to excise the neural tube in the midbrain region. B') Donor embryo with dorsal neural tube graft tissue excised (graft tissue indicated by black arrowheads). Dotted line in diagram indicates where cuts should be made in the quail embryo to excise the donor tissue for unilateral grafting of the midbrain region of the neural tube. C) Chimeric embryo after grafting of quail dorsal neural tube explant into the chick midbrain region.
<img alt="Figure 4" src="/files/ftp_upload/3622/3622fig4.jpg" /> 
Figure 4. Immunofluorescence detecting quail-specific nuclear antigen QCPN in donor-derived cells in the chimeric embryo. A) Whole-mount image of HH11 chimera (grafted at HH9) showing QCPN staining in red, and HNK-1 staining (a marker of migrating NCCs) in green. 10X magnification, dorsal view. B) Cross section through embryo in (A), showing QCPN-positive quail derived cells in the neural tube and migratory NCC co-stained with HNK-1 (green). 40X magnification. C) Cross-section through maxillary process of E5 chimera (incubated for 3 days post graft), showing QCPN-positive NCC-derived cells (red) which have migrated from the grafted region of the neural tube to their final location in the embryo. 10X magnification. D) Section through E5 chimera showing contribution of quail derived NCC to the trigeminal ganglion (red) and differentiation into TuJ1-positive neurons (green). 40X magnification. *, NCCs; MP, maxillary process; NT, neural tube; TG, trigeminal ganglion.

The authors have nothing to disclose.

The grafting of quail neural tube into host chick embryos described here is a straightforward and inexpensive technique for tracing specific subpopulations of migrating NCCs emanating from different regions along the rostro-caudal axis21,67-69. This technique takes advantage of the ease of access to avian embryos (as compared to mammalian embryos) and may be combined with other techniques, such as tissue ablation, injection of inhibitory molecules, or genetic manipulation via electroporation of expression plas...

<table border="1">
<tbody>
 <tr>
 <td>Reagent</td>
 <td>Company</td>
 <td>Catalog number</td>
 </tr>
 <tr>
 <td>Chick eggs</td>
 <td>Various &ndash; we use Texas A&amp;M University's Poultry Science Department, TX.</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Quail eggs</td>
 <td>Various &ndash; we use Ozarks Egg Company, MO.</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Egg incubator (Digital Readout 1502 Sportsman Incubator w/Humidity 110-120 Volt AC)</td>
 <td><a href="http://www.poultrysupply.com" target="_blank">www.poultrysupply.com</a></td>
 <td>1502</td>
 </tr>
 <tr>
 <td>Dumont AA forceps, Inox Epoxy-coated</td>
 <td>Fine Science Tools</td>
 <td>11210-10</td>
 </tr>
 <tr>
 <td>Scotch tape</td>
 <td>Any office supply store</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Curved Iris forceps</td>
 <td>Fine Science Tools</td>
 <td>11065-07</td>
 </tr>
 <tr>
 <td>India ink</td>
 <td>Any art supply store</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Pen/Strep (Penicillin, Streptomycin) Solution </td>
 <td>VWR International</td>
 <td>101447-068</td>
 </tr>
 <tr>
 <td>Clear Packing tape</td>
 <td>Any office supply store</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Needle pulling apparatus</td>
 <td>Narashige, Japan</td>
 <td>PE-21</td>
 </tr>
 <tr>
 <td>Pulled glass needle, made from 1.5-1.8 x 100mm borosilicate glass capillary tube</td>
 <td>Kimble chase</td>
 <td>34500 99</td>
 </tr>
 <tr>
 <td>Pulled glass pipette, made from 5&frac34;" Pasteur pipette</td>
 <td>Fisher Scientific</td>
 <td>13-678-6A</td>
 </tr>
 <tr>
 <td>Mouth pipette apparatus (aspirator tube assembly for calibrated microcapillary pipette)</td>
 <td>Sigma-Aldrich</td>
 <td>A5177-52A</td>
 </tr>
 <tr>
 <td>Dumont #5 forceps</td>
 <td>Fine Science Tools</td>
 <td>11251-30</td>
 </tr>
 <tr>
 <td>Tungsten wire, 0.1mm diameter</td>
 <td>VWR International</td>
 <td>AA10404-H2</td>
 </tr>
 <tr>
 <td>Needle holders (Nickel-plated pin holder)</td>
 <td>Fine Science Tools</td>
 <td>26018-17</td>
 </tr>
 <tr>
 <td>QCPN antiserum</td>
 <td>Developmental Studies Hybridoma Bank, University of Iowa</td>
 <td>QCPN</td>
 </tr>
 <tr>
 <td>Alexa Fluor secondary antibody (e.g., Alexa Fluor 594 goat anti-mouse IgG1)</td>
 <td>Invitrogen</td>
 <td>A21125</td>
 </tr>
 <tr>
 <td>Ringer's Solution (2L):
 <ul>
 <li>14.4g NaCl</li>
 <li>0.34g CaCl2</li>
 <li>0.74g KCl</li>
 <li>0.230g Na2HPO4</li>
 <li>0.04g KH2PO4</li>
 <li>ddH2O to 2L</li>
 <li>Filter and autoclave</li>
 </ul></td>
 <td>All reagents from Fisher Scientific</td>
 <td>
 <ul>
 <li>7647-14-5</li>
 <li>10043-52-4</li>
 <li>7447-40-7</li>
 <li>7558-79-4</li>
 <li>7778-77-0</li>
 </ul></td>
 </tr>
 </tbody>
</table>

analysis of neural crest migration and differentiation by cross-species transplantation

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. Chimeric avian embryos, in which quail donor tissue is transplanted into a chick embryo in ovo, combine the power of indelible genetic labeling of cell populations with the ease of manipulation presented by the avian embryo.
Quail-chick chimeras are a classical tool for tracing migratory neural crest cells (NCCs)1-3. NCCs are a transient migratory population of cells in the embryo, which originate in the dorsal region of the developing neural tube4. They undergo an epithelial to mesenchymal transition and subsequently migrate to other regions of the embryo, where they differentiate into various cell types including cartilage5-13, melanocytes11,14-20, neurons and glia21-32. NCCs are multipotent, and their ultimate fate is influenced by 1) the region of the neural tube in which they originate along the rostro-caudal axis of the embryo11,33-37, 2) signals from neighboring cells as they migrate38-44, and 3) the microenvironment of their ultimate destination within the embryo45,46. Tracing these cells from their point of origin at the neural tube, to their final position and fate within the embryo, provides important insight into the developmental processes that regulate patterning and organogenesis.
Transplantation of complementary regions of donor neural tube (homotopic grafting) or different regions of donor neural tube (heterotopic grafting) can reveal differences in pre-specification of NCCs along the rostro-caudal axis2,47. This technique can be further adapted to transplant a unilateral compartment of the neural tube, such that one side is derived from donor tissue, and the contralateral side remains unperturbed in the host embryo, yielding an internal control within the same sample2,47. It can also be adapted for transplantation of brain segments in later embryos, after HH10, when the anterior neural tube has closed47.
Here we report techniques for generating quail-chick chimeras via neural tube transplantation, which allow for tracing of migratory NCCs derived from a discrete segment of the neural tube. Species-specific labeling of the donor-derived cells with the quail-specific QCPN antibody48-56 allows the researcher to distinguish donor and host cells at the experimental end point. This technique is straightforward, inexpensive, and has many applications, including fate-mapping, cell lineage tracing, and identifying pre-patterning events along the rostro-caudal axis45. Because of the ease of access to the avian embryo, the quail-chick graft technique may be combined with other manipulations, including but not limited to lens ablation40, injection of inhibitory molecules57,58, or genetic manipulation via electroporation of expression plasmids59-61, to identify the response of particular migratory streams of NCCs to perturbations in the embryo's developmental program. Furthermore, this grafting technique may also be used to generate other interspecific chimeric embryos such as quail-duck chimeras to study NCC contribution to craniofacial morphogenesis, or mouse-chick chimeras to combine the power of mouse genetics with the ease of manipulation of the avian embryo.62

Watch this Scientific Journal Video about Analysis of Neural Crest Migration and Differentiation by Cross-species Transplantation at JoVE.com

Analysis of Neural Crest Migration and Differentiation by Cross-species Transplantation

An approach for analyzing migration and eventual fate of avian neural crest cells in quail-chick chimeric embryos is described. This method is a simple and straightforward technique for tracing neural crest cells during migration and differentiation that are otherwise difficult to distinguish within an unmanipulated chick embryo.

Avian embryos provide a unique platform for studying many vertebrate developmental processes, due to the easy access of the embryos within the egg. ...

analysis-neural-crest-migration-differentiation-cross-species

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JoVE Journal

Neuroscience

22.7K 次观看.  Rice University. 描述的分析禽流鹌鹑鸡嵌合胚胎的神经嵴细胞的迁移和最终命运的方法。此方法是一种简单明了的技术追踪神经嵴细胞迁移和分化的过程中，否则难以区分，在未添加鸡胚。

视频: Analysis of Neural Crest Migration and Differentiation by Cross-species Transplantation

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 ...

Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor functions. Most investigations of sensorimotor mechanisms rely on either examination of neurons while an animal is static1,2 or record extracellular neuronal activity during a movement.3,4 While these studies have provided the fundamental background for sensorimotor function, they either do not evaluate functional information which occurs during a movement or are limited in their ability to fully characterize the anatomy, physiology and neurochemical phenotype of the neuron. A technique is shown here which allows extensive characterization of individual neurons during an in vivo movement. This technique can be used not only to study primary afferent neurons but also to characterize motoneurons and sensorimotor interneurons. Initially the response of a single neuron is recorded using electrophysiological methods during various movements of the mandible followed by determination of the receptive field for the neuron. A neuronal tracer is then intracellularly injected into the neuron and the brain is processed so that the neuron can be visualized with light, electron or confocal microscopy (Fig. 1). The detailed morphology of the characterized neuron is then reconstructed so that neuronal morphology can be correlated with the physiological response of the neuron (Figs. 2,3). In this communication important key details and tips for successful implementation of this technique are provided. Valuable additional information can be determined for the neuron under study by combining this method with other techniques. Retrograde neuronal labeling can be used to determine neurons with which the labeled neuron synapses; thus allowing detailed determination of neuronal circuitry. Immunocytochemistry can be combined with this method to examine neurotransmitters within the labeled neuron and to determine the chemical phenotypes of neurons with which the labeled neuron synapses. The labeled neuron can also be processed for electron microscopy to determine the ultrastructural features and microcircuitry of the labeled neuron. Overall this technique is a powerful method to thoroughly characterize neurons during in vivo movement thus allowing substantial insight into the role of the neuron in sensorimotor function.

A technique is described to quantify the in vivo physiological response of mammalian neurons during movement and correlate the physiology of the neuron with neuronal morphology, neurochemical phenotype and synaptic microcircuitry.

The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor ...

Isolation and Culture of Neural Crest Stem Cells from Human Hair Follicles

Hair follicles undergo lifelong growth and hair cycle is a well-controlled process involving stem cell proliferation and quiescence. Hair bulge is a well-characterized niche for adult stem cells1. This segment of the outer root sheath contains a number of different types of stem cells, including epithelial stem cells2, melanocyte stem cells3 and neural crest like stem cells4-7. Hair follicles represent an accessible and rich source for different types of human stem cells. We and others have isolated neural crest stem cells (NCSCs) from human fetal and adult hair follicles4,5. These human stem cells are label-retaining cells and are capable of self-renewal through asymmetric cell division in vitro. They express immature neural crest cell markers but not differentiation markers. Our expression profiling study showed that they share a similar gene expression pattern with murine skin immature neural crest cells. They exhibit clonal multipotency that can give rise to myogenic, melanocytic, and neuronal cell lineages after in vitro clonal single cell culture. Differentiated cells not only acquire lineage-specific markers but also demonstrate appropriate functions in ex vivo conditions. In addition, these NCSCs show differentiation potential toward mesenchymal lineages. Differentiated neuronal cells can persist in mouse brain and retain neuronal differentiation markers. It has been shown that hair follicle derived NCSCs can help nerve regrowth, and they improve motor function in mice transplanted with these stem cells following transecting spinal cord injury8. Furthermore, peripheral nerves have been repaired with stem cell grafts9, and implantation of skin-derived precursor cells adjacent to crushed sciatic nerves has resulted in remyelination10. Therefore, the hair follicle/skin derived NCSCs have already shown promising results for regenerative therapy in preclinical models.
Somatic cell reprogramming to induced pluripotent stem (iPS) cells has shown enormous potential for regenerative medicine. However, there are still many issues with iPS cells, particularly the long term effect of oncogene/virus integration and potential tumorigenicity of pluripotent stem cells have not been adequately addressed. There are still many hurdles to be overcome before iPS cells can be used for regenerative medicine. Whereas the adult stem cells are known to be safe and they have been used clinically for many years, such as bone marrow transplant. Many patients have already benefited from the treatment. Autologous adult stem cells are still preferred cells for transplantation. Therefore, the readily accessible and expandable adult stem cells in human skin/hair follicles are a valuable source for regenerative medicine.

This article presents a robust protocol for isolation and culture of neural crest stem cells from human hair follicles.

Hair follicles undergo lifelong growth and hair cycle is a well-controlled process involving stem cell proliferation and quiescence. Hair bulge is a ...

T-maze Forced Alternation and Left-right Discrimination Tasks for Assessing Working and Reference Memory in Mice

Forced alternation and left-right discrimination tasks using the T-maze have been widely used to assess working and reference memory, respectively, in rodents. In our laboratory, we evaluated the two types of memory in more than 30 strains of genetically engineered mice using the automated version of this apparatus. Here, we present the modified T-maze apparatus operated by a computer with a video-tracking system and our protocols in a movie format. The T-maze apparatus consists of runways partitioned off by sliding doors that can automatically open downward, each with a start box, a T-shaped alley, two boxes with automatic pellet dispensers at one side of the box, and two L-shaped alleys. Each L-shaped alley is connected to the start box so that mice can return to the start box, which excludes the effects of experimenter handling on mouse behavior. This apparatus also has an advantage that in vivo microdialysis, in vivo electrophysiology, and optogenetics techniques can be performed during T-maze performance because the doors are designed to go down into the floor. In this movie article, we describe T-maze tasks using the automated apparatus and the T-maze performance of &alpha;-CaMKII+/- mice, which are reported to show working memory deficits in the eight-arm radial maze task. Our data indicated that &alpha;-CaMKII+/- mice showed a working memory deficit, but no impairment of reference memory, and are consistent with previous findings using the eight-arm radial maze task, which supports the validity of our protocol. In addition, our data indicate that mutants tended to exhibit reversal learning deficits, suggesting that &alpha;-CaMKII deficiency causes reduced behavioral flexibility. Thus, the T-maze test using the modified automatic apparatus is useful for assessing working and reference memory and behavioral flexibility in mice.

This article presents the protocol of T-maze tests using a modified automated apparatus for assessing the learning and memory functions in mice.

Forced alternation and left-right discrimination tasks using the T-maze have been widely used to assess working and reference memory, respectively, in ...

Analysis of Trunk Neural Crest Cell Migration using a Modified Zigmond Chamber Assay

Neural crest cells (NCCs) are a transient population of cells present in vertebrate development that emigrate from the dorsal neural tube (NT) after undergoing an epithelial-mesenchymal transition 1,2. Following EMT, NCCs migrate large distances along stereotypic pathways until they reach their targets. NCCs differentiate into a vast array of cell types including neurons, glia, melanocytes, and chromaffin cells 1-3. The ability of NCCs to reach and recognize their proper target locations is foundational for the appropriate formation of all structures containing trunk NCC-derived components 3. Elucidating the mechanisms of guidance for trunk NCC migration has therefore been a matter of great significance. Numerous molecules have been demonstrated to guide NCC migration 4. For instance, trunk NCCs are known to be repelled by negative guidance cues such as Semaphorin, Ephrin, and Slit ligands 5-8. However, not until recently have any chemoattractants of trunk NCCs been identified 9. 
Conventional in vitro approaches to studying the chemotactic behavior of adherent cells work best with immortalized, homogenously distributed cells, but are more challenging to apply to certain primary stem cell cultures that initially lack a homogenous distribution and rapidly differentiate (such as NCCs). One approach to homogenize the distribution of trunk NCCs for chemotaxis studies is to isolate trunk NCCs from primary NT explant cultures, then lift and replate them to be almost 100% confluent. However, this plating approach requires substantial amounts of time and effort to explant enough cells, is harsh, and distributes trunk NCCs in a dissimilar manner to that found in in vivo conditions. 
Here, we report an in vitro approach that is able to evaluate chemotaxis and other migratory responses of trunk NCCs without requiring a homogenous cell distribution. This technique utilizes time-lapse imaging of primary, unperturbed trunk NCCs inside a modified Zigmond chamber (a standard Zigmond chamber is described elsewhere10). By exposing trunk NCCs at the periphery of the culture to a chemotactant gradient that is perpendicular to their predicted natural directionality, alterations in migratory polarity induced by the applied chemotactant gradient can be detected. This technique is inexpensive, requires the culturing of only two NT explants per replicate treatment, avoids harsh cell lifting (such as trypsinization), leaves trunk NCCs in a more similar distribution to in vivo conditions, cuts down the amount of time between explantation and experimentation (which likely reduces the risk of differentiation), and allows time-lapse evaluation of numerous migratory characteristics.

An approach to analyze the migration of explanted cells (trunk neural crest cells) is described. This method is inexpensive, gentle, and capable of distinguishing chemotaxis from both chemokinesis and other influences on migratory polarity such as those derived from cell-cell interactions within the primary trunk neural crest cell culture.

Neural crest cells (NCCs) are a transient population of cells present in vertebrate development that emigrate from the dorsal neural tube (NT) after ...

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 ...

A Galvanotaxis Assay for Analysis of Neural Precursor Cell Migration Kinetics in an Externally Applied Direct Current Electric Field

The discovery of neural stem and progenitor cells (collectively termed neural precursor cells) (NPCs) in the adult mammalian brain has led to a body of research aimed at utilizing the multipotent and proliferative properties of these cells for the development of neuroregenerative strategies. A critical step for the success of such strategies is the mobilization of NPCs toward a lesion site following exogenous transplantation or to enhance the response of the endogenous precursors that are found in the periventricular region of the CNS. Accordingly, it is essential to understand the mechanisms that promote, guide, and enhance NPC migration. Our work focuses on the utilization of direct current electric fields (dcEFs) to promote and direct NPC migration - a phenomenon known as galvanotaxis. Endogenous physiological electric fields function as critical cues for cell migration during normal development and wound repair. Pharmacological disruption of the trans-neural tube potential in axolotl embryos causes severe developmental malformations1. In the context of wound healing, the rate of repair of wounded cornea is directly correlated with the magnitude of the epithelial wound potential that arises after injury, as shown by pharmacological enhancement or disruption of this dcEF2-3. We have demonstrated that adult subependymal NPCs undergo rapid and directed cathodal migration in vitro when exposed to an externally applied dcEF. In this protocol we describe our lab's techniques for creating a simple and effective galvanotaxis assay for high-resolution, long-term observation of directed cell body translocation (migration) on a single-cell level. This assay would be suitable for investigating the mechanisms that regulate dcEF transduction into cellular motility through the use of transgenic or knockout mice, short interfering RNA, or specific receptor agonists/antagonists.

In this protocol we demonstrate how to construct custom chambers that permit the application of a direct current electric field to enable time-lapse imaging of adult brain derived neural precursor cell translocation during galvanotaxis.

The discovery of neural stem and progenitor  cells (collectively termed neural precursor cells) (NPCs) in the adult  mammalian brain has led to a body of ...

A Quantitative Cell Migration Assay for Murine Enteric Neural Progenitors

Neural crest cells (NCC) are a transient and multipotent cell population that originates from the dorsal neural tube and migrates extensively throughout the developing vertebrate embryo. In addition to providing peripheral glia and neurons, NCC generate melanocytes as well as most of the cranio-facial skeleton. NCC migration and differentiation is controlled by a combination of their axial origin along the neural tube and their exposure to regionally distinct extracellular cues. Such contribution of extracellular ligands is especially evident during the formation of the enteric nervous system (ENS), a complex interconnected network of neural ganglia that locally controls (among other things) gut muscle movement and intestinal motility. Most of the ENS is derived from a small initial pool of NCC that undertake a long journey in order to colonize - in a rostral to caudal fashion - the entire length of the prospective gut. Among several signaling pathways known to influence enteric NCC colonization, GDNF/RET signaling is recognized as the most important. Indeed, spatiotemporally controlled secretion of the RET ligand GDNF by the gut mesenchyme is chiefly responsible for the attraction and guidance of RET-expressing enteric NCC to and within the embryonic gut. Here, we describe an ex vivo cell migration assay, making use of a transgenic mouse line possessing fluorescently labeled NCC, which allows precise quantification of enteric NCC migration potential in the presence of various growth factors, including GDNF.

We present an ex vivo cell migration assay that allows precise quantification of enteric neural crest cell migration potential in the presence of various growth factors.

Neural crest cells (NCC) are a transient and multipotent cell population that originates from the dorsal neural tube and migrates extensively throughout ...