Neural Explant Cultures from Xenopus laevis

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

1. 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).
2. Fertilize eggs in vitro with minced testes, as described previously⁹.
3. 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 °C to slow development.

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.

1. 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/μl, followed by prompt storage at -80 °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.
2. Prepare injection needles by pulling capillary with a needle puller⁹, to a tip diameter of ~ 0.2 μm. 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 example^11,12 ), but we fill the needle with RNA by placing a small drop (0.5 – 1 μl) 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.
3. 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.
4. 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 °C depending upon desired speed of development. (For dissecting neural tubes the following day, use ~ 22 °C. For the day after, use ~ 14 °C.)

3. Neural Tube Dissection and Plating

1. Prior to performing the dissections, prepare the culture dishes¹⁴ . First, coat coverslips with enough solution of 200 μg/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 μg/ml laminin in PBS (we usually use 500 μl 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 μg/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/μl each) to culture media to increase axon outgrowth.
2. 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.
3. 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.
4. 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.
5. 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.
6. Transfer the neural tube to an agarose-coated dish filled with culture media, as described in step 3.1.
7. 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.
8. After plating, do not move the dishes because this would disturb the attaching cells. Incubate the plated neural tube explants around 20-22 °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.

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.

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 μm. Click here to view larger figure.