Neuronal Tracing Using Lipophilic Membrane Dyes Fluorescent Dextrans and Horseradish Peroxidase HRP

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The organization of neurons and their axons in the CNS and peripheral nervous system can be examined using anterograde and retrograde tracing techniques with fluorescent or nonfluorescent dyes. The lipophilic membrane dyes, such as DiI, have the advantage that they may be used on fixed as well as living tissue, whereas the intracellular dyes, such as fluorescent dextrans and HRP, rely to some extent on the mechanisms of axonal transport and thus work best on living tissue. The principle is the same for all the dyes; if you are interested in seeing the neuronal cell bodies, then apply the dyes to their distal axons, and if you are interested in the axonal projections, terminations, or growth cones, then apply the dyes to the neuronal cell bodies. The techniques described below work well on chick embryos up to at least Hamburger and Hamilton stage 25.

3.3.1. Lipophilic Membrane Dyes

1. Fix embryos in 3.5% paraformaldehyde, and store them in this solution in the fridge until you are ready to label them.

2. Dissect in PBS to reveal the part of the CNS or PNS to be labeled. Do not be too rough with the tissue, since this techniques relies on the dye diffusing through intact cell membranes. If you break the membranes it will not work.

3. Back-fill a micropipet (tip opening approx 2-5 pm) with a small quantity of DiI or DiA (D-282 and D-291 from Molecular Probes, respectively). Use a concentration of 3 mg/mL in DMF. DiI fluoresces intensely red and DiA fluoresces also in the red, but more intensely green, and with appropriate filter sets, can readily be used for double-labeling experiments (Fig. 1).

4. Micromanipulate the tip of the micropipet into the tissue, and use a pressure injector (e.g., a Picospritzer II) to deposit the dye. Injections are often made more easily if the pipette is fractionally withdrawn from the full depth of the penetration. If you want to label the cut end of a peripheral nerve, depositing the dye onto the cut surface will efficiently label the axons within.

5. Gently blow away with a Pasteur pipet any excess dye that floats up from the targeted area. If these are allowed to settle onto the embryo, they will label inappropriate areas.

6. Return the specimen to 3.5% paraformaldehyde at room temperature in the dark for 12-48 h.

7. Observe on an epifluorescent microscope either as a whole mount or after sectioning on a cryostat or vibratome. Tissues can be cleared in 90% glycerol in PBS containing 2.5% DABCO (Sigma). The signal in these specimens is not permanent, so they should be analyzed and photographed as soon as possible.

3.3.2. Fluorescent Dextrans

Fluorescent dextrans may in some circumstances have advantages over lipophilic membrane dyes. Those that have a lysine residue can be fixed into the cytoplasm and thus form a nice stable signal. Because the fluorescence is in the cytoplasm they may work better in double-staining procedures, which use cell-surface antibodies. The author finds that for labeling axons within the CNS, dextrans often give more intense staining of neuronal projections than the membrane dyes. For efficient labeling of axons in the PNS, the membrane dyes are, however, the best.

1. Carefully remove embryo from egg and transfer to a Sylgard Dow Corning, Midland, MI)-covered dish containing a physiological saline solution.

2. Free embryo of membranes, and pin it down to dissect and reveal the appropriate area of neural tissue.

3. Mix up a small aliquot of fluorescent dextran. Use Molecular Probes product numbers D-3308 and D-3306 for tetramethylrhodamine and fluorescein fluorescence, respectively. The author keeps a small frozen stock of dextran made up at a concentration of100 mg/mL in distilled water from which he transfer a very small drop onto a Sylgard surface. As the water evaporates off, the dextran becomes sticky and is easily picked up on to the tip of a stainless-steel or tungsten micropin or onto the tips of watchmaker's forceps. Simply add more water to the drop of dextran if it dries out too much.

4. To label axons within the CNS, push a micropin laden with dextran into the region of the axon tract. The dextran is taken up into the axons damaged by the pin, so you can regulate the number of axons labeled by controlling the size of the damage. To label axons in peripheral nerves, the author finds it is more efficient to crush the nerve between the tips of dextran-laden watchmaker's forceps.

5. The cut ends of the axons will seal over in about 30 min to 1 h. Thus, if you wait this long and blow away the excess dextran from the first application, a second application with a differently colored dextran can be made nearby without risking contamination of the first axons.

6. Place the embryo in fresh aerated physiological saline at room temperature for up to 12 h (small embryos will need considerably less time than this, about 3-4 h).

7. Fix tissue in 3.5% paraformaldehyde for at least 2 h, and then observe either as a whole mount or after sectioning. Tissue can be sectioned using a cryostat, vibrotome, or by following conventional wax embedding.

HRP is not a fluorescent tracer (it generates a dense brown reaction product as outlined below), but has the advantage of producing permanent preparations and can be used in combination with whole-mount in situ hybridization techniques.

1.-4. Follow the procedure outlined for labeling with dextrans, but substitute a thick HRP solution for the dextran mixture. For HRP, the author mixes a small pile of crystals with a small drop of distilled water, and wait for it to evaporate to a sticky consistency.

5. Place the embryo in fresh aerated physiological saline at room temperature for up to 12 h (small embryos will need considerably less time than this, about 3-4 h).

6. Fix tissue in 2.5% gluteraldehyde in PBS (pH 7.4) for 1-2 h.

7. Wash thoroughly in several changes of PBS over a period of at least 2 h.

8. Incubate in diaminobenzidine (DAB) (5 mg in 10 mL of PBS) for at least 1 h. DAB is thought to be carcinogenic and should be handled with care. Wear gloves, and use it only in a fume hood.

9. Add hydrogen peroxide to a concentration 0.003% in the DAB solution for about 5-15 min. Carefully look at the reaction in a covered Petri-dish under a dissecting microscope to check its not going too fast. The reaction can be slowed down by reducing the concentration of hydrogen peroxidase. Stop reaction by washing in excess PBS with azide.

10. Discard DAB solutions into an excess of potassium permanganate before disposal.

11. Observe as a whole mount or after sectioning by vibrotome, cryostat, or after wax embedding. Specimens can be cleared in 90% glycerol in PBS or in nonaqueous mountant.

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