The authors would like to thank Bob Freeman for training and the Kirschner lab for the use of the frog facility, and members of the Van Vactor lab for support. We thank the Nikon Imaging Center at Harvard Medical School for assistance with light microscopy for the images in Figure 1. This work was funded by the following: NRSA NIH fellowship and NIH K99 fellowship to L.A.L, Basic Science Partnership funding (<a href="https://bsp.med.harvard.edu/" target="_blank">https://bsp.med.harvard.edu/</a>) to AEF, and NIH RO1 NS035909 to D.V.V.

Note: We describe the steps in the first two sections only in brief, as excellent protocols with detailed information have been published elsewhere that focus more specifically on these steps (e.g.8-12). Additionally, the general protocol of working with Xenopus spinal neurons in live cell culture has been previously published in a detailed methods article8. We highly recommend reviewing that article as a complement to this video, although here we do provide enough information to successfully perform and troubleshoot the neural tube dissection and plating protocol in step 3.
1. In Vitro Fertilization of Xenopus Eggs
<ol>
 <li>Obtain eggs from female frogs that were injected with chorionic gonadotropin (400 units/frog) 12-18 hr before the egg collection. Collect eggs in 1X Marc's Modified Ringer solution (MMR) (0.1 M NaCl, 2.0 mM KCl, 1.0 mM MgSO4, 2.0 mM CaCl2, 5.0 mM HEPES, pH 7.49).</li>
 <li>Fertilize eggs in vitro with minced testes, as described previously9.</li>
 <li>After at least 20 min, remove embryo jelly coat by incubating embryos in 2% cysteine in 1X MMR (brought to pH 7.8 with NaOH) for 3-5 min. Wash with 0.1X MMR (or 0.1X MBS (Modified Barth's Saline, 1X recipe: 88 mM NaCl, 1 mM KCl, 0.7 mM CaCl2, 1 mM MgSO4, 2.5 mM NaHCO3, 5 mM HEPES, pH 7.8)) 3-5 times, and keep embryos at room temperature until injections, or if desired, place embryos at 14-18 &deg;C to slow development.</li>
</ol>
2. Microinjection of RNA
Note: while we utilize RNA injection here, these techniques are not limited to RNA, and DNA, proteins, or modified nucleic acids for gene manipulation can also be used and have been described previously12.
<ol>
 <li>Prior to experiment, RNAs for labeling cytoskeletal or other structures are transcribed from linearized DNA templates using the mMessage mMachine kit (Ambion). mRNAs require a 5' cap and a 3' polyadenylation tail to promote translation and prevent degradation in embryos. pCS2+ is a commonly-used vector for this purpose, and the transcription kit includes a cap analog during the synthesis reaction. We re-suspend transcribed mRNA in nuclease-free ddH20, at a stock concentration between 500 and 2,000 ng/&mu;l, followed by prompt storage at -80 &deg;C. Prior to injection, dilute RNA in ddH20 to the appropriate concentration for injection. As only a small volume will be injected into embryos, resuspending in buffer is not necessary. Our final injection concentration is typically 50-200 pg/nl, depending on the construct, and we usually will inject up to 4 nl per embryo, although many labs traditionally inject up to 10 nl, which is approximately 1% of the total volume of an embryo. RNA can be toxic at high doses, and the dose per embryo generally ranges from 10 pg to 1 ng, although up to 5 ng can be tolerated depending on the particular gene and purity. However, for imaging protein localization, the lowest possible level should be injected to avoid over-expression artifacts. RNA for EB1-GFP in this protocol is used at final amount of 250 pg per embryo.</li>
 <li>Prepare injection needles by pulling capillary with a needle puller9, to a tip diameter of ~ 0.2 &mu;m. With a Sutter P-87 puller, we use the following settings: heat 644, pull 125, velocity 70, time 250, but these parameters differ depending on machine and must be re-adjusted after changing the filament (we currently use a 2.5 mm square box filament that is 2.5 mm wide). Under a microscope, break needle tip with forceps at an angle to generate a quill-like shape. There are other methods for breaking and filling injection needles (for example11,12 ), but we fill the needle with RNA by placing a small drop (0.5 - 1 &mu;l) to the back end of the needle. For microinjecting into embryos, there are a number of commercially available injection systems, two of the most common being the Pico-injector (Medical Systems), and the Nanoject (Drummond Scientific) (see11 for more information). We use the Medical Systems PLI-100 Pico-Injector, which uses compressed gas for a digitally set period of time in order to deliver consistent nanoliter volumes. Injection volume must be calibrated for each new micropipette using a stage micrometer, and injection time should be adjusted to achieve desired amount of injected RNA.</li>
 <li>Place fertilized embryos at the 1- to 4-cell stage into a plastic dish containing 5% Ficoll in 0.1X MMR. Holding the embryo in place with forceps, or placing in a holding platform (a plastic mesh, with a ~1 mm grid, adhered to the bottom of the dish with plasticine or dental wax), inject desired volume into animal blastomeres (see step 2.1 for details regarding injection volumes). Distribute in several locations throughout embryo, at least one injection into each of the animal blastomeres, to obtain more uniform distribution (for example, for a 4-cell stage embryo, we typically inject 1-2 times in each blastomere, whereas in a 2-cell stage embryo, we inject 2-4 times in each blastomere). A benefit of waiting until the 2-4 cell stage is that you ensure the embryo has begun to cleave normally. However, if time is of the essence, you can start at the 1-cell stage, with little difference in the end result other than more embryos needing to be injected. Older stage embryos can be used as well, and this is particularly useful if specific cell types are targeted for injection, but as we generally prefer more broad expression, we inject in younger embryos to reduce the need for more numerous injections.</li>
 <li>Transfer injected embryos to a plastic dish containing 0.1X MMR and allow them to develop until stage 20-2313. Incubate embryos at 14 to 22 &deg;C depending upon desired speed of development. (For dissecting neural tubes the following day, use ~ 22 &deg;C. For the day after, use ~ 14 &deg;C.)</li>
</ol>
3. Neural Tube Dissection and Plating
<ol>
 <li>Prior to performing the dissections, prepare the culture dishes14 . First, coat coverslips with enough solution of 200 &mu;g/ml poly-lysine in PBS to pool over the surface of the coverslip, of either Mattek or Lab-Tek culture dishes, and incubate for one hour (alternatively, one could use glass coverslips sitting loose in a Petri dish, for later placement on glass slides for imaging and immunocytochemistry). Aspirate and wash with an excess of PBS 3 times, and let dry. Then coat dishes with 10 &mu;g/ml laminin in PBS (we usually use 500 &mu;l per coverslip, enough to pool over the surface) for one hour at 37 degrees. Aspirate laminin solution and wash 3 times with an excess of PBS, being careful not to let laminin-coated surfaces become exposed to the air interface. Replace PBS with culture media (50% Ringer's, 49% L-15 media, 1% Fetal Bovine Serum, plus 50 &mu;g/ml penicillin/streptomycin and gentamycin, pH 7.4 and filter sterilized). (Ringer's Media, 115 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 10 mM Hepes pH 7.4, 0.5 mM EDTA). While our protocol uses this media, it should be noted that this is an enriched media common for older Xenopus retinal ganglion cultures, where neurons have reduced energy reserved. Young spinal cord cultures will survive and grow for more than 24 hr in pure Ringers's Media without additional growth factors, and this is indeed one of the benefits of using this system. Optional: add NT3 and BDNF (to final concentrations of 25 ng/&mu;l each) to culture media to increase axon outgrowth.</li>
 <li>Because of the variability in the expression of injected mRNA, embryos may exhibit a mosaic of fluorescence. Prior to performing dissections, screen embryos for the presence of fluorescence, to identify those embryos which express the fluorescent fusion protein in the neural tube. Place fluorescent embryos at stage 20-23 into an agarose-coated plastic dish filled with Steinberg's media (58 mM NaCl, 0.67 mM KCl, 0.44 mM Ca(NO3)2, 1.3 mM MgSO4, 4.6 mM Tris, pH to 7.8 then autoclaved)5. To make the agarose-coated dish, coat bottom of plastic dish with melted 1% agarose in 0.1X MMR and let harden. This provides a soft surface so that damage does not occur to the fine forceps and tungsten needles used during the subsequent dissection steps. Dissection of embryos beyond stage 23 is possible, but it is more challenging as tissues adhere more closely to each other and thus require longer treatment with collagenase.</li>
 <li>Under a dissecting scope, remove the vitelline membrane with fine forceps, and then isolate the entire dorsal portion of the embryo, by making a series of incisions. While one forceps hold the embryo in place, use the second to make an incision on the side of the embryo to expose the hollow interior. Then, use both forceps to pinch along the tissue between the dorsal and ventral halves of the embryo, thereby cutting the embryo in half to isolate the dorsal portion containing the neural tube.</li>
 <li>Place the dorsal explant in an Eppendorf tube with 2 mg/ml collagenase in Steinberg's media for 15-20 min on a rotator, to loosen tissues, and then pipette dorsal explant into a plastic agarose-coated dish with fresh Steinberg's solution. Alternatively, the explant can be placed in a Petri dish containing the collagenase solution for 15 min, where the entire dissection could then be carried out.</li>
 <li>Using a pair of forceps, gently dissect the neural tube from the dorsal epidermis and the ventral notochord. Slide the tip of the forceps between the epidermis and underlying tissue and slowly pull back the epidermis, revealing the neural tube beneath. Then, use one forceps to hold the tissue and another to slide the tip between the neural tube and notochord. Finally, use forceps to remove the somites on either side of the tube.</li>
 <li>Transfer the neural tube to an agarose-coated dish filled with culture media, as described in step 3.1.</li>
 <li>After collection of several neural tubes, cut them into numerous pieces (approximately 20, if the whole tube is isolated) using sharpened tungsten needles or forceps, and then plate the pieces in culture dishes filled with medium (prepared in step 3.1), spreading out the explants evenly in rows.</li>
 <li>After plating, do not move the dishes because this would disturb the attaching cells. Incubate the plated neural tube explants around 20-22 &deg;C. We typically leave the dish of cells on the bench at room temperature overnight. One of the benefits of Xenopus laevis neurons is that a special incubator for cell culture that regulates CO2 levels or temperature is not needed. Neurites and growth cones can be observed and imaged at room temperature 12-24 hr after plating, although outgrowth can be accelerated with the addition of growth factors. In general, expression of a RNA or plasmid product will result in variable expression between cells, and thus there may be a range of fluorescence expression levels between growth cones. We have observed RNA expression of fluorescent proteins to persist beyond 48 hr after plating, although this depends on the particular construct.</li>
</ol>
4. Representative Results
After following the protocol, which is summarized in Figure 1A, healthy, correctly-dissected and cultured neural explants will send out numerous axons or neurites in every direction on laminin/poly-lysine substrate, shown in Figure 1B. The growth cones at the tips of the axons can be imaged by either DIC optics, as seen in Figure 1C, to image overall motility dynamics, or high-resolution fluorescence microscopy to examine the localization of fluorescently-tagged cytoskeletal proteins, for example, EB1-GFP is shown in Figure 1D.
<img alt="Figure 1" src="/files/ftp_upload/4232/4232fig1.jpg" /> 
Figure 1. Summary of experimental protocol and expected outcome. A) Protocol flow-chart. See movie for details regarding dissection on day 3. The entire neural tube should be cut into about 20 equally-sized pieces and spread out evenly in rows on the coverslip. In this cartoon, scissors are depicted to represent cutting the tissue with fine forceps. CG chorionic gonadotropin B) DIC image of axons or neurites growing out of explant (explant at top left corner). C) Higher magnification image of growth cone at tip of a growing axon. D) Fluorescence microscopy image of EB1-GFP in growth cone. Scale bar 10 &mu;m. <a href="https://www.jove.com/files/ftp_upload/4232/4232fig1large.jpg" target="_blank"> Click here to view larger figure</a>.

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	<li>Lowery, L. A., Van Vactor, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+trip+of+the+tip:+understanding+the+growth+cone+machinery.">The trip of the tip: understanding the growth cone machinery.</a> Nat. Rev. Mol. Cell Biol. 10, 332-343 (2009).</li><li>Buck, K. B., Zheng, J. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Growth+cone+turning+induced+by+direct+local+modification+of+microtubule+dynamics.">Growth cone turning induced by direct local modification of microtubule dynamics.</a> J. Neurosci. 22, 9358-9367 (2002).</li><li>Lee, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+microtubule+plus+end+tracking+protein+Orbit/MAST/CLASP+acts+downstream+of+the+tyrosine+kinase+Abl+in+mediating+axon+guidance.">The microtubule plus end tracking protein Orbit/MAST/CLASP acts downstream of the tyrosine kinase Abl in mediating axon guidance.</a> Neuron. 42, 913-926 (2004).</li><li>Santiago-Medina, M., Myers, J. P., Gomez, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+adhesion+and+signaling+dynamics+in+Xenopus+laevis+growth+cones.">Imaging adhesion and signaling dynamics in Xenopus laevis growth cones.</a> Dev. Neurobiol. , (2011).</li><li>Tanaka, E. M., Kirschner, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubule+behavior+in+the+growth+cones+of+living+neurons+during+axon+elongation.">Microtubule behavior in the growth cones of living neurons during axon elongation.</a> J. Cell Biol. 115, 345-363 (1991).</li><li>Tanaka, E., Kirschner, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+microtubules+in+growth+cone+turning+at+substrate+boundaries.">The role of microtubules in growth cone turning at substrate boundaries.</a> J. Cell Biol. 128, 127-137 (1995).</li><li>Tanaka, E., Ho, T., Kirschner, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+microtubule+dynamics+in+growth+cone+motility+and+axonal+growth.">The role of microtubule dynamics in growth cone motility and axonal growth.</a> J. Cell Biol. 128, 139-155 (1995).</li><li>Gomez, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Working+with+Xenopus+spinal+neurons+in+live+cell+culture.">Working with Xenopus spinal neurons in live cell culture.</a> Methods Cell Biol. 71, 129-156 (2003).</li><li>Sive, H. L., Grainger, R. M., Harland, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Early Development of Xenopus laevis: A Laboratory Manual 2010. , (2010).</li><li>Cross, M. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+of+the+action+potential+mechanism+of+amphibian+neurons+isolated+in+culture.">The development of the action potential mechanism of amphibian neurons isolated in culture.</a> Proc. Natl. Acad. Sci. U.S.A. 73, 1641-1645 (1976).</li><li>Turney, S. G., Bridgman, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laminin+stimulates+and+guides+axonal+outgrowth+via+growth+cone+myosin+II+activity.">Laminin stimulates and guides axonal outgrowth via growth cone myosin II activity.</a> Nat. 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Biol. 10, 849-852 (2000).</li></ol>

No conflicts of interest declared.

Xenopus laevis neural explants send out neurites in a very robust manner by 24 hr after plating on the laminin/poly-lysine substrate if conditions are appropriate. With this substrate, growth cones are highly motile and can achieve axon lengths of up to 1 mm, extending in all directions outward from the explant, although typical lengths are 100 &mu;m or more. If neurites do not grow out, there are a limited number of reasons for this to be the case. One possibility is that the neural explants did not properly a...

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 </tr>
 <tr>
 <td>Chorionic Gonadotropin</td>
 <td>Argent Chemical Laboratories</td>
 <td>C-HCG-ON-10</td>
 </tr>
 <tr>
 <td>Cysteine</td>
 <td>Sigma-Aldrich</td>
 <td>52-90-4</td>
 </tr>
 <tr>
 <td>mMessage mMachine kit</td>
 <td>Ambion</td>
 <td>AM1340</td>
 </tr>
 <tr>
 <td>Capillary Borosil Needles 1.2 mm (OD) x 0.9 mm (ID)</td>
 <td>FHC</td>
 <td>30-31-0</td>
 </tr>
 <tr>
 <td>Ficoll</td>
 <td>Sigma</td>
 <td>F2637</td>
 </tr>
 <tr>
 <td>Dumont #5 Biologie Inox Forceps</td>
 <td>Fine Science Tools</td>
 <td>11252-20</td>
 </tr>
 <tr>
 <td>Collagenase</td>
 <td>&nbsp;Sigma-Aldrich</td>
 <td>&nbsp;9001-12-1</td>
 </tr>
 <tr>
 <td>Mattek dishes</td>
 <td>Mat Tek Corporation</td>
 <td>P35G-1.5-14-C</td>
 </tr>
 <tr>
 <td>L-15</td>
 <td>Invitrogen</td>
 <td>21083-027</td>
 </tr>
 <tr>
 <td>Poly-l-lysine</td>
 <td>Sigma</td>
 <td>P-1399</td>
 </tr>
 <tr>
 <td>Laminin</td>
 <td>Sigma</td>
 <td>L2020</td>
 </tr>
 <tr>
 <td>NT3</td>
 <td>Sigma</td>
 <td>N1905</td>
 </tr>
 <tr>
 <td>BDNF</td>
 <td>Sigma</td>
 <td>B3795</td>
 </tr>
 </tbody>
</table>

neural explant cultures from xenopus laevis

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During axon outgrowth, the growth cone must integrate multiple sources of guidance cue information to modulate its cytoskeleton in order to propel the growth cone forward and accurately navigate to find its specific targets1. How this integration occurs at the cytoskeletal level is still emerging, and examination of cytoskeletal protein and effector dynamics within the growth cone can allow the elucidation of these mechanisms. Xenopus laevis growth cones are large enough (10-30 microns in diameter) to perform high-resolution live imaging of cytoskeletal dynamics (e.g.2-4 ) and are easy to isolate and manipulate in a lab setting compared to other vertebrates. The frog is a classic model system for developmental neurobiology studies, and important early insights into growth cone microtubule dynamics were initially found using this system5-7 . In this method8, eggs are collected and fertilized in vitro, injected with RNA encoding fluorescently tagged cytoskeletal fusion proteins or other constructs to manipulate gene expression, and then allowed to develop to the neural tube stage. Neural tubes are isolated by dissection and then are cultured, and growth cones on outgrowing neurites are imaged. In this article, we describe how to perform this method, the goal of which is to culture Xenopus laevis growth cones for subsequent high-resolution image analysis. While we provide the example of +TIP fusion protein EB1-GFP, this method can be applied to any number of proteins to elucidate their behaviors within the growth cone.

Watch this Scientific Journal Video about Neural Explant Cultures from Xenopus laevis at JoVE.com

Neural Explant Cultures from Xenopus laevis

Culturing neural explants from dissected Xenopus laevis embryos that express fluorescent fusion proteins allows for imaging of growth cone cytoskeletal dynamics.

Neural Explant Cultures from Xenopus laevis

The complex process of axon guidance is largely driven by the growth cone, which is the dynamic motile structure at the tip of the growing axon. During ...

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Neuroscience

12.5K Views.  Harvard Medical School. Культивирование нейронных эксплантов от расчлененного Xenopus Хепориз Эмбрионов, которые выражают флуоресцентных белков слияния позволяет изображений конуса роста динамики цитоскелета.

Video: Neural Explant Cultures from Xenopus laevis

Derivation of Enriched Oligodendrocyte Cultures and Oligodendrocyte/Neuron Myelinating Co-cultures from Post-natal Murine Tissues

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications for understanding the pathogenesis of demyelinating diseases such as Multiple Sclerosis (MS). Cellular development is commonly studied with primary cell culture models. Primary cell culture facilitates the evaluation of a given cell type by providing a controlled environment, free of the extraneous variables that are present in vivo. While OL cultures derived from rats have provided a vast amount of insight into OL biology, similar efforts at establishing OL cultures from mice has been met with major obstacles. Developing methods to culture murine primary OLs is imperative in order to take advantage of the available transgenic mouse lines. 
Multiple methods for extraction of OPCs from rodent tissue have been described, ranging from neurosphere derivation, differential adhesion purification and immunopurification 1-3. While many methods offer success, most require extensive culture times and/or costly equipment/reagents. To circumvent this, purifying OPCs from murine tissue with an adaptation of the method originally described by McCarthy &amp; 
de Vellis 2 is preferred. This method involves physically separating OPCs from a mixed glial culture derived from neonatal rodent cortices. The result is a purified OPC population that can be differentiated into an OL-enriched culture. This approach is appealing due to its relatively short culture time and the unnecessary requirement for growth factors or immunopanning antibodies. 
While exploring the mechanisms of OL development in a purified culture is informative, it does not provide the most physiologically relevant environment for assessing myelin sheath formation. Co-culturing OLs with neurons would lend insight into the molecular underpinnings regulating OL-mediated myelination of axons. For many OL/neuron co-culture studies, dorsal root ganglion neurons (DRGNs) have proven to be the neuron type of choice. They are ideal for co-culture with OLs due to their ease of extraction, minimal amount of contaminating cells, and formation of dense neurite beds. While studies using rat/mouse myelinating xenocultures have been published 4-6, a method for the derivation of such OL/DRGN myelinating co-cultures from post-natal murine tissue has not been described. Here we present detailed methods on how to effectively produce such cultures, along with examples of expected results. These methods are useful for addressing questions relevant to OL development/myelinating function, and are useful tools in the field of neuroscience.

This article describes methods to derive enriched populations of murine oligodendrocyte precursor cells (OPCs) in primary culture, which differentiate to produce mature oligodendrocytes (OLs). In addition, this report describes techniques to produce murine myelinating co-cultures by seeding mouse OPCs onto a neurite bed of mouse dorsal root ganglion neurons (DRGNs).

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications ...

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 ...

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

The process by which the anterior region of the neural plate gives rise to the vertebrate retina continues to be a major focus of both clinical and basic research. In addition to the obvious medical relevance for understanding and treating retinal disease, the development of the vertebrate retina continues to serve as an important and elegant model system for understanding neuronal cell type determination and differentiation1-16. The neural retina consists of six discrete cell types (ganglion, amacrine, horizontal, photoreceptors, bipolar cells, and M&uuml;ller glial cells) arranged in stereotypical layers, a pattern that is largely conserved among all vertebrates 12,14-18.
While studying the retina in the intact developing embryo is clearly required for understanding how this complex organ develops from a protrusion of the forebrain into a layered structure, there are many questions that benefit from employing approaches using primary cell culture of presumptive retinal cells 7,19-23. For example, analyzing cells from tissues removed and dissociated at different stages allows one to discern the state of specification of individual cells at different developmental stages, that is, the fate of the cells in the absence of interactions with neighboring tissues 8,19-22,24-33. Primary cell culture also allows the investigator to treat the culture with specific reagents and analyze the results on a single cell level 5,8,21,24,27-30,33-39. Xenopus laevis, a classic model system for the study of early neural development 19,27,29,31-32,40-42, serves as a particularly suitable system for retinal primary cell culture 10,38,43-45.
Presumptive retinal tissue is accessible from the earliest stages of development, immediately following neural induction 25,38,43. In addition, given that each cell in the embryo contains a supply of yolk, retinal cells can be cultured in a very simple defined media consisting of a buffered salt solution, thus removing the confounding effects of incubation or other sera-based products 10,24,44-45.
However, the isolation of the retinal tissue from surrounding tissues and the subsequent processing is challenging. Here, we present a method for the dissection and dissociation of retinal cells in Xenopus laevis that will be used to prepare primary cell cultures that will, in turn, be analyzed for calcium activity and gene expression at the resolution of single cells. While the topic presented in this paper is the analysis of spontaneous calcium transients, the technique is broadly applicable to a wide array of research questions and approaches (Figure 1).

Dissection, Culture, and Analysis of Xenopus laevis Embryonic Retinal Tissue

Xenopus laevis provides an ideal model system for studying cell fate specification and physiological function of individual retinal cells in primary cell culture. Here we present a technique for dissecting retinal tissues and generating primary cell cultures that are imaged for calcium activity and analyzed by in situ hybridization.

The process by which  the anterior region of the neural plate gives rise to the vertebrate retina  continues to be a major focus of both clinical and ...

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

Much information about the coupling of presynaptic ionic currents with the release of neurotransmitter has been obtained from invertebrate preparations, most notably the squid giant synapse1. However, except for the preparation described here, few vertebrate preparations exist in which it is possible to make simultaneous measurements of neurotransmitter release and presynaptic ionic currents. Embryonic Xenopus motoneurons and muscle cells can be grown together in simple culture medium at room temperature; they will form functional synapses within twelve to twenty-four hours, and can be used to study nerve and muscle cell development and synaptic interactions for several days (until overgrowth occurs). Some advantages of these co-cultures over other vertebrate preparations include the simplicity of preparation, the ability to maintain the cultures and work at room temperature, and the ready accessibility of the synapses formed2-4. The preparation has been used widely to study the biophysical properties of presynaptic ion channels and the regulation of transmitter release5-8. In addition, the preparation has lent itself to other uses including the study of neurite outgrowth and synaptogenesis9-12, molecular mechanisms of neurotransmitter release13-15, the role of diffusible messengers in neuromodulation16,17, and in vitro synaptic plasticity18-19.

Simultaneous Pre- and Post-synaptic Electrophysiological Recording from Xenopus Nerve-muscle Co-cultures

This video demonstrates the procedures used to grow primary cultures of embryonic Xenopus nerve and muscle cells and the usefulness of this preparation for making simultaneous pre- and post-synaptic patch clamp recordings.

Much information about the coupling of  presynaptic ionic currents with the release of neurotransmitter has been obtained  from invertebrate preparations, ...

One Mouse, Two Cultures: Isolation and Culture of Adult Neural Stem Cells from the Two Neurogenic Zones of Individual Mice

The neurosphere assay and the adherent monolayer culture system are valuable tools to determine the potential (proliferation or differentiation) of adult neural stem cells in vitro. These assays can be used to compare the precursor potential of cells isolated from genetically different or differentially treated animals&#160;to determine the effects of exogenous factors on neural precursor cell proliferation and differentiation and to generate neural precursor cell lines that can be assayed over continuous passages. The neurosphere assay is traditionally used for the post-hoc identification of stem cells, primarily due to the lack of definitive markers with which they can be isolated from primary tissue&#160;and has the major advantage of giving a quick estimate of precursor cell numbers in brain tissue derived from individual animals. Adherent monolayer cultures, in contrast, are not traditionally used to compare proliferation between individual animals, as each culture is generally initiated from the combined tissue of between 5-8 animals. However, they have the major advantage that, unlike neurospheres, they consist of a mostly homogeneous population of precursor cells and are useful for following the differentiation process in single cells. Here, we describe, in detail, the generation of neurosphere cultures and, for the first time, adherent cultures from individual animals. This has many important implications including paired analysis of proliferation and/or differentiation potential in both the subventricular zone (SVZ) and dentate gyrus (DG) of treated or genetically different mouse lines, as well as a significant reduction in animal usage.

Here we describe a detailed protocol for the simultaneous generation of neural precursor cell cultures, as either adherent monolayers or neurospheres, from the subventricular zone and dentate gyrus of individual adult mice.

The neurosphere assay and the adherent monolayer culture system are valuable tools to determine the potential (proliferation or differentiation) of adult ...

A Simple Behavioral Assay for Testing Visual Function in Xenopus laevis

Measurement of the visual function in the tadpoles of the frog, Xenopus laevis, allows screening for blindness in live animals. The optokinetic response is a vision-based, reflexive behavior that has been observed in all vertebrates tested. Tadpole eyes are small so the tail flip response was used as alternative measure, which requires a trained technician to record the subtle response. We developed an alternative behavior assay based on the fact that tadpoles prefer to swim on the white side of a tank when placed in a tank with both black and white sides. The assay presented here is an inexpensive, simple alternative that creates a response that is easily measured. The setup consists of a tripod, webcam and nested testing tanks, readily available in most Xenopus laboratories. This article includes a movie showing the behavior of tadpoles, before and after severing the optic nerve. In order to test the function of one eye, we also include representative results of a tadpole in which each eye underwent retinal axotomy on consecutive days. Future studies could develop an automated version of this assay for testing the vision of many tadpoles at once.

A Simple Behavioral Assay for Testing Visual Function in Xenopus laevis

Xenopus laevis tadpoles prefer swimming on the white side of a black/white tank. This behavior is guided by their vision. Based on this behavior, we present a simple assay to test the visual function of tadpoles.

Measurement of the visual function in the tadpoles of the frog, Xenopus laevis, allows screening for blindness in live animals. The optokinetic response ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...

Derivation of Leptomeninges Explant Cultures from Postmortem Human Brain Donors

Even though great progress has been made in the clinical characterization of Parkinson&#39;s disease, several studies report that the diagnosis of Parkinson&#39;s disease is not pathologically confirmed in up to 25% of clinically diagnosed Parkinson&#39;s disease. Therefore, tissue collected from clinically diagnosed patients with idiopathic Parkinson&#39;s disease can have a high rate of misdiagnosis; hence in vitro studies from such tissues to study Parkinson&#39;s disease as a preclinical model can become futile.
By collecting postmortem human leptomeninges with a confirmed neuropathological diagnosis of Parkinson&#39;s disease and characterized by nigrostriatal cell loss and intracellular protein inclusions called Lewy bodies, one can be certain that clinically observed parkinsonism is not caused by another underlying disease process (e.g. tumor, arteriosclerosis).
This protocol presents the dissection and preparation of postmortem human leptomeninges for derivation of a meningeal fibroblast culture. This procedure is robust and has a high success rate. The challenge of the culture is sterility as the brain procurement is generally not performed under sterile conditions. Therefore, it is important to supplement the culture media with a cocktail of penicillin, streptomycin, and amphotericin B.
The derivation of meningeal fibroblasts from autopsy-confirmed cases with Parkinson&#39;s disease provides the foundation for in vitro modeling of Parkinson&#39;s disease. Meningeal fibroblasts appear 3-9 days after sample preparation and about 20-30 million cells can be cryopreserved in 6-8 weeks. The meningeal fibroblast culture is homogenous and the cells express fibronectin, a commonly used marker to identify meninges.

The leptomeninges explant culture protocol from human postmortem brain is a technically robust and simple way to derive fibronectin-positive meningeal fibroblasts within 6-8 weeks and cryopreserve approximately 20-30 million cells.

Even though great progress has been made in the clinical characterization of Parkinson&#39;s disease, several studies report that the diagnosis of ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...

Simultaneous Recordings of Cortical Local Field Potentials, Electrocardiogram, Electromyogram, and Breathing Rhythm from a Freely Moving Rat

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls body functions and internal organ rhythms when animals are exposed to emotional challenges and changes in their living environments. In general experiments, signals from different organs, such as the brain and the heart, are recorded by independent recording systems that require multiple recording devices and different procedures for processing the data files. This study describes a new method that can simultaneously monitor electrical biosignals, including tens of local field potentials in multiple brain regions, electrocardiograms that represent the cardiac rhythm, electromyograms that represent awake/sleep-related muscle contraction, and breathing signals, in a freely moving rat. The recording configuration of this method is based on a conventional micro-drive array for cortical local field potential recordings in which tens of electrodes are accommodated, and the signals obtained from these electrodes are integrated into a single electrical board mounted on the animal's head. Here, this recording system was improved so that signals from the peripheral organs are also transferred to an electrical interface board. In a single surgery, electrodes are first separately implanted into the appropriate body parts and the target brain areas. The open ends of all of these electrodes are then soldered to individual channels of the electrical board above the animal's head so that all of the signals can be integrated into the single electrical board. Connecting this board to a recording device allows for the collection of all of the signals into a single device, which reduces experimental costs and simplifies data processing, because all data can be handled in the same data file. This technique will aid the understanding of the neurophysiological correlates of the associations between central and peripheral organs.

This study introduces a method for the simultaneous recording of local field potentials in the brain, electrocardiograms, electromyograms, and breathing signals of a freely moving rat. This technique, which reduces experimental costs and simplifies data analysis, will contribute to the understanding of the interactions between the brain and peripheral organs.

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls ...