Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

The overall goal of this procedure is to record the synaptic and excitability properties of embryonic zebra fish mouth their cells. This is accomplished by first dissecting the zebra fish embryo to expose the ventral surface of the hind brainin. The second step is to place the dissected preparation onto the stage of a patch clamp microscope, and identify the mouth in their cells.

Next patch. Clamp onto the M cells using the appropriate recording mode and record synaptic activity and excitability properties. The final step is to analyze the recorded data to ascertain synaptic or excitability properties.

Ultimately, patch clamping from these cells is used to show the properties of synaptic receptors in a developing organism. Generally, individuals new to this method will have two main areas of difficulty. The first is the dissection.

The second formation of the gig scene can be challenging. I first had the idea for this method when I decided to work on a zebrafish preparation in which the hind brain was attached to the spinal cord, demonstrating the procedure will be burram Roy, a PhD graduate student in my lab, Begin this procedure by preparing the dissecting dish lined with Sard. First, fix the washers to the slide with grease.

Then add liquid sard to the center of the washer. The sard usually cures overnight and forms a small circular bed on the glass slide, which becomes the base of the dissecting dish. After placing a cover slip on the slide, place the slide into the recording chamber to serve as the base for the dissection.

Afterward, prepare the required solutions. The extracellular solutions should be left at room temperature while intracellular solutions should be kept sterile. One can also add 0.1%loose for yellow to the intracellular solution, so that visualization of M cell morphology may be performed at the end of the experiment to confirm the cell identity.

Next, raise the wild type zebra fish embryos in egg water at 28.5 degrees Celsius and collect and stage according to the previous publication. For dissection, transfer the embryos to the dissecting dish that also serves as the recording chamber. Then anesthetize them for seven to 10 minutes.

In an anesthetizing salt solution that closely approximates physiological saline, one can determine if they're adequately anesthetized by examining the response to a gentle tail pinch. Afterward, pin the embryos through the no accord and onto the SIL guard line dish with 0.001 inch tungsten wire under a dissecting microscope with 25 x magnification. Adjust the magnification on the dissecting scope to 40 to 50 x to perform the dissection.

Now, remove the jaw eyes and forebrain using a fine pair of forceps. Gently peel the hindbrain away from the underlying node cord by carefully working the forceps between the node cord and the brain. Then gently turn the hind brain over so that the ventral surface faces up.

This positioning provides good access to the M cells, which are usually within five to 10 micrometers of the ventral surface in young embryos and 20 to 50 micrometers in older larva. Next, carefully move the preparation to the recording setup where the patch clamp experiment can be performed. The M cells are visualized with differential interference contrast, although other modes of imaging may also be used to prepare for patch clamping.

Begin by pulling some patch CLA pipettes with thin-walled bo silicate glass. In a horizontal pull pipette tip diameters are about 0.2 to 0.4 micrometers after fire polishing to a smooth edge, and the shank taper is about four millimeters long. Fill the pipette tip with intracellular solution by dipping the tip into the intracellular fluid.

Then insert a syringe needle into the pipette and gently expel intracellular solution from the syringe. To finish filling the pipette, attach the pipette to the amplifier head stage In the electrophysiology setup, it is important to keep the head stage at a roughly 45 degree angle to the horizontal axis as this ensures an entry angle for the pipette that is suitable for the formation of high resistant seals with the mouth in their cell right before the pipette lowers into the bath solution, apply a small amount of positive pressure to the pipette to reduce the chance of the tip blocking. Continue to approach the M cell with a small amount of positive pressure in the pipette.

The positive pressure gently pushes the cell from side to side, and when positioned immediately over the cell forms a small dimple on the cell membrane. Now leave the pipette in place for a few seconds to gently clean the cell surface so that a strong seal between the pipette and the membrane can be formed. Then release the positive pressure in the pipette to initiate the seal.

A small amount of negative pressure coupled with negative pipette potential results in giga seals forming within a few seconds. Subsequently, change the holding potential in the amplifier to minus 60 millivolts. Rupture the cell membrane with a series of short pulses of suction.

Then immediately record membrane potentials and minimize capacitance artifacts. Compensate the cell capacitance and access resistance by 70 to 85%Access resistance should be monitored every 30 seconds to a minute, and if there's a change of 20%or more, abort the experiment. Once the experiment has ended and enough data has been acquired, sacrifice the embryo by removing the hind brain with a pair of forceps.

This figure shows the representative recordings of excitatory A MPA and NMDA receptor currents obtained from 48 HPF embryos. Here are the spontaneous A MPA ME PSCs recorded from an M cell at a holding potential of minus 60 millivolts. And this is the average ME PSCs.

Here are the spontaneous A MPA and NMDA me PSCs recorded from an M cell at a holding potential of 40 millivolts. And this is the average ME PSCs glutamate me PSCs were recorded in the presence of one micromolar TTX to block action potentials five micromolar strict nine and 100 micromolar ritoxin to block glycine and G occurrence shown. Here are the spontaneous MI PSCs recorded from an M cell at a holding potential of minus 60 millivolts.

And the average MI PSCs MI PSCs were recorded in the presence of one micromolar TTX to block action potentials. One millimolar chimeric acid to block glutamate receptor activity and 10 micromolar by Qing to block GABA occurrence. These are the action potentials recorded from an M cell in this particular cell.

A SRA threshold stimulus of 0.48 nano amps elicited several action potentials, but a stimulus to threshold only results in the production of a single action potential. Once mastered, this technique can be done in two to four hours if performed properly. After watching this video, you should have a good understanding of how to patch clamp record from M cells and other reticular spinal neurons in zebrafish embryos.