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

The overall goal of this procedure is to obtain primary cultures of synaptically coupled spinal neurons and muscle cells from xenopus embryos. This is accomplished by first inducing breeding in xenopus by injection of human chorionic in atropin into both male and female frogs and collecting fertilized eggs. The second step is to remove the jelly coat and fatel and membranes from stage 22 embryos.

Next, spinal neurons and muscles are isolated from the embryos and the skin is removed and discarded. The final step is to plate the isolated cells into culture dishes and allow the formation of functional neuromuscular junctions. Ultimately, functional synaptic connections can be demonstrated with simultaneous paired patch clamp recordings from neurons and muscle cells in culture.

The beauty of this technique is that it yields a readily accessible vertebrate synaptic preparation. It can be used to monitor neurotransmitter release with postsynaptic electrophysiological recordings while simultaneously measuring presynaptic ion occurrence. Furthermore, the methods are clear and straightforward, requiring no special equipment beyond a dissecting microscope, and the so cultures can then be stored at room temperature in a simple culturing medium.

This preparation can be useful in answering key questions in the field of synaptic physiology, including how pre-synaptic, biochemical, and biophysical events are coupled with neurotransmitter release. In addition, it is also useful for the study of neuromodulation synaptogenesis and synaptic plasticity To begin this procedure, prepare the required solutions according to the accompanying manuscript. Then fabricate at least two micro dissection tools by gluing a minuchin pin to the end of a glass pastier pipette.

With Sano acrylic glue, allow the sharp end of the pin to extend past the end of the pipette approximately 0.5 centimeters. Two days before preparing the cultures, identify a breeding ready pair of XUS by observing a prominent reddish claw aika on the female and dark pigmentation on the planar surface of the front paws of the male. Next net each frog and hold it ventral side down in a sink with the net so that it can escape.

Inject one milliliter of HCG through the net and subcutaneously into one of the dorsal lymph sacks to ensure that the animals will nott trample the newly laid and fertilized eggs. Install a screened floor with a mesh size of half inch fitted one to two inches at the bottom of the tank. Then place the breeding pair together in a covered 10 gallon tank of water.

Leave the frogs undisturbed for 12 to 48 hours until fertilized eggs are observed on the floor below the screen. Then remove the animals from the tank, but leave the eggs undisturbed for at least 24 hours longer. This pair of frogs can be rebred after six weeks.

Next, loosen the embryos from the bottom of the tank and transfer them to four or five 60 by 15 millimeter culture dishes containing 10%saline. Sort the embryos by stage according to the scheme of new coupe and Faber embryos that are smooth in appearance with light, brown and white modeling like those on the left are ideal. On the other hand, embryos that have large black or white patches like those on the right are generally unhealthy and unusable.

Inside a laminar flow hood label and fill approximately halfway 360 by 15 millimeter sterile culture dishes with 10%saline and one with CMF. Using a sterile glass pastier pipette transfer five to 10, stage 22 to 24 embryos into one of the dishes containing 10%saline with the eight of a stereo zoom dissecting microscope inside the hood, remove the jelly coat and viel membrane from each embryo using two pairs of sterile forceps. 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.

Next, hold each embryo gently but firmly with a pair of forceps and use the micro dissection tool to remove the neural tube and the 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. Then move each dissected neural tube myotome to a clean part of the dish away from the yolk granules and other debris.

After about 15 minutes in CMF, use the forceps to lift the pigmented skin free of the dissected tissue and discard the skin. After an additional 30 to 60 minutes, the cells will form a sand pile as they become dissociated from one another. In this procedure, thaw a 10 milliliter tube of culture medium, next at 70 microliters of ITS in 35 nanograms per milliliter, BDNF label and fill the 35 millimeter sterile culture dishes approximately halfway with L 15 culture medium.

Use one culture dish for each embryo you have dissected. To fabricate a plating pipette, hold both ends of a glass pastier pipette and place the tapered portion over a flame. Then pull the ends apart to about 10 centimeters.

Subsequently break off the end to yield to tip of approximately 0.2 millimeters. Afterward, use the plating pipette to suck the sand pile up from one embryo. Be careful to minimize the amount of solution drawn.

Next, expel the cells onto the bottom of a culture dish and plate them into several lines. Then leave the dishes of plated cells undisturbed for at least 15 minutes to allow the cells time to attach to the dish. After 12 to 24 hours in culture, plated cells will develop distinct morphological characteristics.

For example, muscle cells become spindle shaped and spinal neurons develop long processes. Functional synaptic contact between urate, varicosities and muscle cells can be confirmed with simultaneous paired electrophysiological recordings. To do so, prepare two patch electrodes for recording, one for the presynaptic varicosity, and one for the muscle cell.

Then fill the presynaptic electrode with info terin containing solution and the post-synaptic electrode with potassium internal solution. After that, replace the media in the culture dish with NFR. Then transfer it to an inverted microscope fitted with face contrast optics, patch the presynaptic varicosity with the amphotericin containing electrode and the post-synaptic muscle cell with the other electrode to confirm the functionality of the synapse, depolarize the varicosity with a patch clamp amplifier, and observe the simultaneous presynaptic and postsynaptic currents.

This figure shows a five x power view of cells and culture immediately after plating, and this is the same culture at 40 x. Here is the neuromuscular junction in culture, which shows the neuronal soma, presynaptic, varicosity, and postsynaptic muscle cell. Shown here are the presynaptic and postsynaptic currents.

In response to the application of 10 millivolt incremental voltage steps presented to the presynaptic varicosity from minus 30 millivolts to plus 40 millivolts, the holding potential for both cells was minus 70 millivolts Once mastered, this technique can be completed in about two hours. Viable cell cultures can then be obtained in as soon as 24 hours and can be maintained for several days. This preparation has already been in use for a number of years, helping to explain some of the fundamental properties of neuromuscular transmission.

It's our hope that this video will allow for the continuation of discovery by helping investigators become more familiar with this powerful and simple technique.