Channelrhodopsin2 Mediated Stimulation of Synaptic Potentials at Drosophila Neuromuscular Junctions

This procedure begins with growing drosophila. Third, instar maggots with the appropriate genotype on food containing all trans retinol. Each maggot is pinned out head and tail.

Then FLA flat. The brain and posterior nerves are then cut to stop rhythmic motor activity. Next, the blue LED light source with control box is adjusted and focused on the preparation.

Finally, use a sharp electrode to impale muscle six. Then deliver light pulses with a voltage source. Hi, I'm Nicholas Marstein from the lab of Dr.Leslie Griffith, the Department of Biology at Brandeis University.

Hi, I'm Stefan Pulver, also from the Griffith Lab. Today we will show you procedure for evoking synaptic potentials with channel root absent two at the neuromuscular junction of drosophila larva. We use this procedure in our lab to study synaptic physiology and drosophila.

We also use it to teach students synaptic physiology and teaching labs. So let's get started. To get started with this protocol, maintain UAS channel redsin two and okay, 3 7, 1 gal four fly lines in separate bottles containing standard fly media.

Collect virgins from the, okay, 3 7, 1 gal, four fly lines and males from the UAS channel redsin. Two lines. Next, make fly media that is mixed with one millimolar, all trans retinol or a TR.To do this melt regular fly media in a microwave for approximately one minute.

Once melted, allow the media to cool for about 30 to 60 seconds, and then add 100 microliters of 100 millimolar a TR in 100%ethanol for every 10 milliliters of fly media. When the vials containing a TR media are ready, put the collected males and females in each vial. Place the vials in a dark area at 22 to 25 degrees Celsius to let the flies mate and lay eggs wait four to five days until third in star larva are visible.

At this point, you can dispose of the adult flies. We are now ready to proceed with the electrophysiology rig set up. Attach any dissecting scope eyepiece to a blue LED light source with a heat sink.

Use a post and clamp to attach a magnetic base to the eyepiece and LED light source. Place the magnetic base on the air table of the electrophysiology rig. Next, connect the light source to a control circuit, and then connect the control circuit to an external voltage source.

We use the voltage output from the power lab electrophysiology data acquisition system made by ad instruments. Once connected to the voltage source, give one to five volt pulses to the control circuit in order to activate the blue light, adjust the magnetic base and the light source until the blue light beam is focused on the area to be occupied by the larval dissection. After making the proper adjustments, we are ready to move on to the larval dissection.

To get started with the dissection, place six 0.1 millimeter pins into the floor of a sogar lined dish. Remove a third in star larvae from the food media and place it into any plastic Petri dish. Rinse the larvae with saline to remove any food that is attached to the larvae.

Then placed the larvae in the dissecting dish near the pulled pins and filled the dish halfway with saline. Now orient the larvae so that you can see two silvery tubes running along the animal's dorsal surface. These silvery tubes are the trachea.

Insert a large pin directly into the tail in between the tracheal tubes. Hold the larvae down and place a second large pin into its head. Be sure to stretch the body out lengthwise as the pin is inserted.

Use scissors to make a small incision near the tail. Continue the incision up the length of the body. Make sure that the tips of the scissors are raised to avoid cutting the ventral nerves and or body wall muscles.

Place four pins on the four corners of the animals. Now open midsection. Set the pins into the dissecting dish so as to filet the animal.

Also pin the preparation so that it's taught using forceps and scissors. Remove the animal's, digestive organs, trachea, and fat bodies. Rinse the prep with saline.

After rinsing, locate the frontal lobes and ventral ganglion of the prep using micro scissors. Carefully cut through the ventral ganglion just behind the frontal lobes. Once the dissection is complete, proceed with the blue light stimulation and muscle recording.

To perform the muscle recording, pull a 10 to 20 mega ohm resistance electrode. Using an electronic electrode puller, fill the pulled electrode with three molar KCL. Place the filled electrode in the electrode holder.

Now place the larval prep under a dissecting microscope on the rig. Adjust the blue light so that the dissected prep is centered in the light beam. Identify muscle six or M six in any body wall segment of the larva using a micro manipulator, carefully insert electrode into M six and watch for rapid hyperpolarization muscle.

Resting membrane potentials should be anywhere from negative 30 millivolts to negative 70 millivolts. Once the electrode is properly inserted, give 20 to 100 millisecond voltage pulses to the control circuit. Incrementally increase the voltage supplied to the control circuit.

Watch for exci junction potentials in the muscle cell. Here is a representative result that shows an exci junction potential evoked by short light pulses. The exci junction potential amplitude shown here is the summed amplitude from two motor neurons that are both known to ate.

M six. Lower intensities or shorter durations of light can expose the smaller amplitude excitatory junction potential from a single motor neuron. We've just shown you how to perform channel road absent two mediated stimulation of synaptic potentials at the oph neuromuscular junction.

When doing this procedure, it's important to remember not to damage the body wall of the prep during dissection. It's also important to remember that although a prep may be overexcited just following a dissection, it will quiet down over time and allow for good recordings. When impaling muscles, be careful not to go below the first layer of muscles into the body wall.

When your electrode is stuck into the body wall. You can sometimes obtain hyperpolarized potentials but not actually be in a muscle cell. So that's it.

Thanks for watching and good luck with your experiments.