100 ng/mL

3.1.2. Preparing the Tissue Culture Dishes for Plating Neurons

Different tissue culture dishes can be used to grow sympathetic neurons. Some protocols require glass cover slips, and others use Aclar dishes. Our protocol uses the 35-mm standard plastic tissue culture dishes from Falcon. The 60-mm, 6-well or multiwell tissue culture dishes can also be used in place of the 35-mm dish.

1. Dissolve poly-dl-ornithine in borate buffer at a concentration of 500 pg/mL. Filter-sterilize, and store aliquots at -20°C.

2. One day prior to dissection, add 1 mL of poly-dl-ornithine into each 35-mm tissue culture dish. Place dishes in a 37°C CO2 incubator overnight (see Note 6).

3. Four to 5 h prior to dissection, remove poly-dl-ornithine from all the dishes, and wash each dish three times with 1 mL of tissue culture-grade water. After the final wash, add 1 mL of laminin (10 pg/mL diluted in CMF saline G buffer) to each dish, and place them back in the incubator. Allow laminin to coat the surface of the dishes for 4-5 h before starting the dissection.

3.2. Culturing Dissociated Rat Embryonic Sympathetic Ganglia

All subsequent protocols including the dissection are carried out in a laminar flow hood to minimize contamination.

3.2.1. Dissection of Embryonic Rat Sympathetic Ganglia

1. Sterilize dissection tools by autoclaving and then immersing them in 70% ethanol.

2. Prepare a pregnant Sprague-Dawley female rat (E15.5) for dissection. Euthanize the rat by placing it in a CO2-filled chamber for 5 min. Pinch the rat's hind footpad and observe for reflex to verify that the rat has expired.

3. Place the rat on a styrofoam surgical stage layered with absorbent diapers, with her back facing down, and then immobilize the rat by pinning the hind legs down to the

Fig. 1. Location of the superior cervical ganglia (SCG) within the thin slice of the lower mandible.

surgical stage with pins. Next, sterilize the fur around her underbelly by dousing the fur with 70% ethanol.

4. Using the tissue forceps, lift skin and make an incision 2-3 cm ventral to the urogenital opening with the operating scissors. Make a ventral midline incision all the way to the diaphragm. Do not cut through the muscle layer yet; just separate the skin from the muscle tissue.

5. Next, make a second incision at the muscle tissue at the location of the first incision near the urogenital opening. As described in step 3, open the peritoneal cavity by cutting through the muscle tissue from the urogenital opening to diaphragm. Avoid lacerating any organs within the peritoneal cavity.

6. Using a sterile tissue forceps, gently lift the embryos from each horn of the uterus. Detach the embryos from the tissue by cutting off the fallopian tube. Transfer the string of embryos to a tissue culture dish.

7. Using a pair of sterile microdissecting tweezers, carefully remove each embryo from the uterine sac. Avoid decapitating the embryos during the extraction, as they are fragile. Make sure the placenta, as well as the amniotic membrane, has been removed from each embryo. Transfer the embryos to a tissue culture dish filled with Sal G buffer. Wash the embryos gently by swirling the dish and buffer around. Repeat wash one more time by transferring the embryos to a new dish. Do not discard the buffer from the first embryo wash (see Note 7).

8. After both washes, transfer the embryos to a glass Petri dish bottom containing Sal G buffer. With the aid of a dissection microscope, decapitate the embryos at the neckline using a pair of sterile microdissecting knives (Fig. 1). Next, make an incision just below the ear to obtain a thin slice of the lower mandible (dotted lines in Fig. 1; see Note 8). Repeat for all embryos.

Fig. 2. A magnified thin-slice image of the lower mandible. The solid ellipse outlines the tongue, and the dotted circles denote the location of the SCG. The dotted midline incision from the larynx to the spinal column separates the lower half of the thin slice for easier access to the SCG.

9. Transfer all the thinly sliced lower mandibles to a new glass Petri dish bottom containing Sal G buffer. Before starting the microdissection, align each slice so that the larynx and tongue are facing up and on top, and the spinal column is at the bottom (Fig. 2). With the microdissecting knives, first remove the spinal column. Next, make an incision in the center of each slice from the larynx to where the spinal column was located (dotted line in Fig. 2). This incision separates the bottom half of the slice, revealing the ganglia. Finally, gently tease apart the surrounding tissue to extract the ganglia from both sides of the slice (circles in Fig. 2; see Note 9).

Fig. 3. A magnified image of the superior cervical ganglion (2) that is still attached to the carotid artery. The nodose ganglion (1) is usually adjacent to the SCG.

10. Each superior cervical ganglion (SCG) is attached to the carotid artery and is tightly associated with the neighboring nodose ganglion (Fig. 3). Since they are adjacent to each other, problems may arise when attempting to differentiate the two ganglia. The SCG is crescent-shaped, and the carotid artery is often still attached to the ganglion when first extracted. The nodose ganglion, which contains sensory neurons and is attached to the vagus nerve, looks like a chicken drumstick and has a short stem.

11. Dissect the remaining slices, and extract all the ganglia. A typical dissection should yield 10 embryos with 20 ganglia. Sequester all the dissected ganglia in one corner of the dish free of any discarded tissue debris. This action will help maintain a cleaner and more homogenous culture.

3.2.2. Plating the Dissociated Sympathetic Neuron Culture

1. Obtain a glass Pasteur pipet with a flame-smoothened tip, and coat the inner surface of the pipet with the first embryo wash buffer (see Subheading 3.2.1., step 7).

This solution prevents the dissected ganglia from adhering to the surface of the glass pipet (see Note 10).

2. Using the glass Pasteur pipet, gently transfer the dissected ganglia from the glass Petri dish bottom into a 15-mL screw-cap conical tube. Avoid pipeting too much liquid into the tube. Briefly centrifuge the suspension in a clinical centrifuge to pellet the ganglia.

3. Remove the supernatant from the conical tube with a micropipet, and add 1 mL of CMF saline G buffer. Allow the tube to sit in the flow hood for 5 min before pelleting the ganglia in a clinical centrifuge.

4. Carefully remove the CMF saline G buffer with a micropipet, and add 1 mL of trypsin (250 |g/mL). Make sure that all the ganglia are covered in trypsin and they remain settled at the bottom of the tube.

5. Place the tube in a 37°C water bath and incubate for 15 min.

6. During this incubation period, wash the laminin-coated tissue culture dishes twice with CMF saline G buffer. After the final wash, remove all excess liquid from the dishes, and allow the dishes to dry in the flow hood (see Note 11).

7. After trypsinization, remove the tube from the water bath, and spin for 1 min in a clinical centrifuge to pellet the ganglia. At this stage, the trypsinized ganglia should be "sticky" and clumped together. Carefully remove the trypsin using a micropipet and add 1 mL of trypsin inhibitor (1 mg/mL). Place the tube in a 37°C water bath for another 5 min.

8. Pellet the ganglia by spinning for 1 min in a clinical centrifuge. Remove the trypsin inhibitor with a micropipet and add 2 mL of the neuron culture medium.

9. Using a glass Pasteur pipet prepared as described in step 1, gently triturate the trypsinized ganglia by repeatedly pipeting up and down. The ganglia will slowly shrink and dissociate into individual neurons. Triturate until most of the ganglia are no longer visible by naked eye (see Note 12).

10. Place the tube in the flow hood for 1 min to allow the undissociated ganglia to settle at the bottom of the tube.

11. Using a micropipet, pipet 100 |L of the dissociated neuron culture onto a laminin-coated tissue culture dish. Before plating, make sure the dish is dry. Place the drop of culture in the center of the dish. The surface tension should hold the drop together and prevent the neurons from flowing to the sides of the dish. Repeat this procedure for all 20 tissue culture dishes.

12. Add 1.9 mL of the neuron culture medium to each dish by dribbling the medium down the sides of the dish and placing the dishes back in the 37°C CO2 incubator (see Note 13).

3.2.3. Maintaining the Dissociated Sympathetic Neuron Culture

1. Two days after dissection, the neuron cell bodies will be attached to the substratum at the center of the dish. Some of the cell bodies will already have neurites. To eliminate the dividing nonneuronal cells, add 1 ||M; Ara-C. Remove Ara-C treatment the next day by replacing with fresh neuron culture medium. Repeat Ara-C treatment, if necessary, 4 or 5 d after the dissection.

2. Typically, neuron culture medium is replaced once every 3 d. Allow the fresh medium to warm up and equilibrate in a 37°C CO2 incubator for at least 30 min prior to replacing the medium (see Note 14).

3. Neurons are generally cultured for 2-3 wk to allow for terminal differentiation before they are ready for viral infection.

3.3. Growth and Differentiation of PC12 Cells With Nerve Growth Factor

PC12 cells are derived from a rat pheochromocytoma cell line that differentiates into sympathetic-like neurons in response to NGF (4). These cells have a doubling time of approx 48 h and do not adhere well to either plastic or glass substrates, which necessitates coating dishes with either a positively charged molecule or an extracellular matrix component. The major advantages of using a cell line, compared with primary tissue, is that variability from experiment to experiment is reduced, large amounts of cells can be grown, and animals are not needed. To maximize this benefit, we routinely differentiate a large number of PC12 cells and store them "primed" ready for use (5). For a comprehensive discussion of the methods for routine culture and other experimental protocols with PC12 cells, see ref. 6.

We routinely grow PC12 cells on plastic dishes coated with collagen, according to the original method of culturing these cells, although poly-D-lysine appears to work equally well for attachment. However, growing PC12 cells on a glass surface requires coating the glass with a combination of poly-ornithine and laminin. The different substrates are described just below.

3.3.1. Collagen Substrate

Coat the inside surface of plastic dishes with collagen at 5 pg/cm2 in 0.02 N acetic acid as follows (see Note 15):

1. Aseptically apply the appropriate volume of collagen to each dish for 1 h at room temperature (Table 2) (see Note 16).

2. Aspirate unbound collagen and rinse dish three times with appropriate volume of sterile water.

3. Dishes are ready for use or they can be stored at 4°C for 1 wk.

3.3.2. Poly-dl-Ornithine Substrate (see Note 17)

1. Dissolve entire 100 mg bottle of poly-dl-ornithine in 200 mL of 0.1 M borate buffer, and filter-sterilize.

3. Aseptically coat dishes using 0.5 mg/mL of poly-dl-ornithine using volumes listed in Table 2.

4. Incubate dishes overnight in a humidified incubator at 37°C.

5. Rinse dishes five times with sterile water using volumes in Table 2.

Table 2

Volumes of Substrates and Washes for Preparing Coated Dishes

Table 2

Volumes of Substrates and Washes for Preparing Coated Dishes

Dish diameter

Volume of substrate applied

dH20 wash


(mL; see Note 16)


0 0

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