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

1. Pre-experimental Preparation

1. Prepare the following solutions (see Table 1 for compositions): (a) 1 liter NFR (Normal Frog Ringer), (b) 100 ml 10% saline, (c) 100 ml CMF (Ca²⁺ /Mg²⁺-free solution), (d) 1 ml ITS (Insulin-Transferrin-Selenium, Sigma I1884), (e) 100 ml L-15 culture medium, (f) 10 ml HCG (human chorionic gonadotropin Sigma CG-10), (g) 100 ml K⁺-internal solution, (h) 100 ml K⁺-internal solution for Amphotericin B. The osmotic strength of solution 1 should be checked to ensure that it is approximately 260 mOsM using a vapor pressure osmometer (e.g. Wescor model #5100C).
2. Filter sterilize solutions (b) and (c). Solutions (a) (b) and (c) can be stored up to three months at 4 °C. Prepare solution (d) by adding 1 ml deionized H₂O to the vial containing the lyophilized powder via a 25 gauge syringe needle and syringe filter. This final solution can be stored at 4 °C for 30 days.
3. Prepare solution (e) in a laminar flow hood by combining all components in a beaker or flask. Filter sterilize the final solution and transfer 10 ml aliquots into sterile centrifuge tubes. Store at -20 °C.
4. Prepare the HCG (solution (f)) by adding 10 ml deionized H₂0 to the vial containing the lyophilized powder via a 25 gauge syringe needle and syringe filter. This final solution can be stored at 4 °C for 30 days.
5. Fabricate at least two microdissection tools by gluing a Minutien pin (26002-10, Fine Science Tools) to the end of a glass Pasteur pipette with cyanoacrylate glue. Allow the sharp end of the pin to extend past the end of the pipette approximately 0.5 cm.
6. Two days before preparing cultures, induce breeding with the following procedure. Identify a breeding-ready pair of Xenopus by observing a prominent, reddish cloaca on the female and dark pigmentation on the planar surface of the front paws of the male. One at a time, net each frog and hold it ventral side down on a sink with the net so that it cannot escape. Inject 1 ml of solution (f) through the net and subcutaneously into one of the dorsal lymph sacs.
7. Place the breeding pair together in a covered ten gallon tank of water. To ensure that the animals will not trample the newly laid and fertilized eggs, install a screened floor with a mesh size of approximately ½” fitted approximately 1-2 inches above the bottom of the tank (Figure 1A).
8. Leave frogs undisturbed for 12-48 hr until the animals are in amplexus and fertilized eggs are observed on the floor below the screen (Figure 1B).
9. Remove the animals from the tank, but leave the eggs undisturbed for at least 24 hr longer. This pair of frogs can be re-bred after six weeks.
10. Loosen the embryos from the bottom of the tank and transfer them to four or five 60×15 mm culture dishes containing 10% saline (solution b). Sort the embryos by stage according to the scheme of Niewkoop and Faber (ref. 20). Useful embryos will be those at stage 22-24. Figure 2A shows an embryo at approximately stage 22 while Figure 2B shows an embryo that is too far along in development to be useful (approximately stage 28). It is important to choose embryos that are healthy: those that are smooth in appearance with light brown and white mottling are ideal. Embryos that have large black or white patches are generally unhealthy and unusable.

2. Microdissection of Xenopus Embryos

1. Inside a laminar flow hood, label and fill approximately halfway three 60×15 mm sterile culture dishes with 10 % saline; and one with CMF.
2. Using a sterile, glass Pasteur pipette transfer five to ten stage 22-24 embryos into one of the dishes containing 10% saline. With the aid of a stereo zoom dissecting microscope inside the hood (0.6-5x with 10 x eyepieces), remove the jelly coat and vitelline membrane from each embryo using two pairs of sterile #5 forceps (11251-30, Fine Science Tools). (See Figures 3A and 3B.)
3. Wash the bare embryos by passing them, one at a time, through the remaining two dishes of 10% saline and finally into the dish containing CMF. Use a new, sterile pipette for each transfer and minimize the volume of solution transferred from dish to dish.
4. In turn, hold each embryo gently but firmly with a pair of forceps and, using the microdissection tool fashioned in step 1.5, remove the neural tube and associated myotomes which are located at the most dorsal aspect of the animal. Do this by making three slices, one at either end of the location of the neural tube and a third just ventral to it (Figure 4A). Move each dissected neural tube/myotome to a clean part of the dish away from the yolk granules and other debris (Figure 4B). The dorsal-ventral axis, and the locations of the neural tube and myotomes are indicated in Figure 4.
5. After about fifteen minutes in this solution (CMF) use forceps to lift the pigmented skin free of the dissected tissue and discard. After an additional 30-60 min, the cells will form a “sand pile” as they become dissociated from one another (Figure 4C).

3. Preparation of Nerve-muscle Co-cultures

1. Thaw a 10 ml tube of culture medium and aseptically add 70 μl ITS and 35 ng/ml BDNF (Sigma B3795).
2. Label and fill sterile 35 mm culture dishes (FD35-100 World Precision Instruments) approximately halfway with L-15 culture medium (solution (e)), one for each embryo dissected.
3. Fabricate a plating pipette by grasping each end of a glass Pasteur pipette while holding the tapered portion over a flame. Pull the ends apart to about 10 cm and then break off the end to yield a tip of approximately 0.2 mm.
4. Use the plating pipette to suck up the “sand pile” from one embryo minimizing the amount of solution drawn. Expel the cells onto the bottom of a culture dish. Plate the cells into several lines. Observe plated undifferentiated cells (Figure 5A and 5B).
5. Leave the dishes of plated cells undisturbed for at least fifteen minutes to allow the cells time to attach to the dish.

4. Patch Clamping Nerve-muscle Synapses

1. After 12-24 hr in culture, plated cells take on distinct morphological characteristics: muscle cells become spindle-shaped and spinal neurons extend long processes (Figure 6). Functional synaptic contact between neurite varicosities and muscle cells can be confirmed with simultaneous paired electrophysiological recordings. “M”, “S” and “V” refer respectively to the muscle cell, neuronal soma and presynaptic varicosity.
2. Prepare two patch electrodes for recording: one for the presynaptic varicosity and one for the muscle cell.
3. For the presynaptic electrode, prepare an amphotericin-containing solution as follows: Add 100 μl DMSO to a 1.5 ml microcentrifuge tube containing 5 mg amphotericin B (Sigma A4888). Vortex this solution for 10 sec. Next, add 10 μl of this solution to a second microcentrifuge tube that contains 625 μl K⁺-internal solution for Amphotericin B (solution (h)). Vortex as above.
4. Fill the pre-synaptic electrode with the amphotericin-containing K⁺-internal solution prepared in step 4.3 and the post-synaptic electrode with K⁺-internal solution (solution (g)).
5. Replace media in the culture dish with NFR and relocate it to an inverted microscope fitted with phase contrast optics.
6. Position both electrodes just above their respective targets. Obtain the whole-cell configuration with the muscle cell electrode before the perforated patch configuration with the varicosity electrode.
7. To confirm functionality of the synapse, depolarize the varicosity with a patch clamp amplifier (e.g. Axopatch 200B, Molecular Devices) under software control (e.g. by pCLAMP 9, Molecular Devices) and observe the simultaneous presynaptic and postsynaptic currents. Figure 7 shows the currents seen in response to depolarizing steps of voltage given to the presynaptic varicosity in 10 mV increments from -30 mV to +40 mV. The inward currents seen in the presynaptic cell are carried by Na⁺ and Ca²⁺ and the outward currents by K⁺. Currents recorded in the postsynaptic cell are responses to neurotransmitter released from the varicosity and are carried mostly by Na⁺ through nicotinic acetylcholine receptor channels.