Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this procedure is to record the single unit responses in vivo from an identified population of neurons in a non-genetic model system. This is accomplished by first performing a craniotomy to expose the brain region of interest. The second step is to use tungsten wires to place the fluorescent dye in the region to which the target neurons project.

Next, the dye is retrograde transported, resulting in selective labeling of the projection neurons. The final step is to record from the labeled axon of the individual projection neuron. Ultimately, single unit extracellular recordings are used to show how central sensory neurons in an intact circuit respond to the sensory stimulation in evo.

This method can help to answer key questions regarding how neuro circuits detect sub millisecond differences in the timing of synaptic inputs. The main advantage of this technique over existing methods is the ability to target individual neurons or even axons and non-genetically accessible systems. Weekly electric fish use electric signals for communication and navigation.

They distinguish signals from different fish using sub millisecond differences in the timing of synaptic inputs. We asked how the presumed time comparator neurons in their electros sensory circuit are able to detect such small timing differences. But this technique can also be applied to other systems such as visual and auditory sensory pathways.

It is particularly difficult to liquid from this cells because of the small size dense miring in the surrounding tissue and the presynaptic terminals that covers most of the cell belly Intracellular dye injection during a whole cell recording. In vitro labels a single cell to guide targeted recordings from that axon. Similarly, retrograde transport following extracellular dye injection provides a means of selectively labeling the projection neurons in superficial brain regions.

This can be used to provide visual guidance for in vivo single unit recordings. To prepare the D coded needle first electrolytic sharpen a 160 micrometer diameter tungsten wire to the final needle tip diameter of five to 50 micrometers. Then the night before the experiment code the needle tip with a two millimolar solution of 10, 000 molecular weight dextran conjugated Alexa floor dye to induce general anesthesia.

Place the fish in tank water with MS 2 22 solution at 300 milligrams per liter immobilize and electrically silence the fish by injecting 100 microliters of flail at three milligrams per milliliter into the dorsal body musculature. Next, fill the recording chamber with tank water and place the fish ventral side down on the platform in the center of the chamber. Then deliver the aerated MS 2 22 solution at 100 milligrams per liter in one to two milliliters per minute by placing a pipette tip in its mouth.

Afterwards, stabilize the fish with fixed rods and wax on both sides of the body. Monitor the fish's health condition by checking the continuous blood flow in the ocular vessels and a normal body color. Then rotate the platform along the vertical axis.

Adjust one end of the platform so that one side of the dorsal surface of the fish's head is exposed above the water while the rest of the fish's body remains submerged. Place a small piece of Kim wipe on any nons submerged portion of the skin to prevent drying. In this procedure, apply 0.4%lidocaine to the exposed surface of the head using a Q-tip.

Then use a scalpel blade to cut a rectangular piece of skin tissue of approximately three millimeters by five millimeters for a 6.2 centimeter fish. The lateral edge of the rectangle should align with the center of the eye. The anterior edge of the rectangle should be just posterior to the eye, and the medial edge of the rectangle should be just lateral to the midline.

Subsequently, remove the rectangle using a pair of forceps. Cut an additional 2.5 millimeter square anterior mely to expose the skull. Completely clear the exposed surface of the skull by scraping away any excess tissue using the scalpel blade.

Then completely dry the surface with a Kim wipe and forced air. Next, glue a metal post to the anterior medial exposed skull region with super glue. Wait until the glue is completely dry.

Use a dental drill with a 0.5 millimeter diameter ball mill carbide tip to draw a rectangle of approximately two millimeters by four millimeters on the skull for a 6.2 centimeter fish. After that, cut the perimeter of the rectangle with the scalpel and forceps. Then peel the skull away.

To expose the brain, cut both the pigmented dura mater and the clear pia mater using a pair of spring scissors or a needle. Then remove the cut portions with a pair of forceps to expose the anterior extra lateral nucleus and the posterior extra lateral nucleus. Now position a manipulator with a D coated needle prepared earlier above the target region containing the axons of interest.

The posterior extra lateral nucleus in our case swiftly insert the needle approximately 25 micrometers into the tissue. After the dye has diffused for 15 to 30 seconds, retract the needle. Repeat with additional fresh needles as needed.

Place each one in a different location so that the dye is distributed throughout the target region. Then rinse the excess dye from the cavity with hickman's ringer solution and wait for at least two hours for dye uptake and transport. To visualize the axons of interest, place the recording chamber with the fish underneath the objective of an upright fixed stage epi fluorescence microscope.

Switch the respiration solution to fresh tank water and maintain the same flow rate. Then place one end of the ground wire in the exposed brain cavity and connect the other end to the ground of the recording head stage. Next, place a pair of recording electrodes next to the base of the tail.

Then connect the electrodes to a differential amplifier and recording device. In order to monitor the electric organ discharge command or EODC, prepare a scaled sketch of the brain region viewed at low magnification, including major blood vessels as landmarks to identify the exact location of labeled axons. To confirm D placement first, view the entire tissue with brightfield illumination for orientation.

Then view it with fluorescent illumination. The posterior extra lateral nucleus should show diffuse labeling. After that, under high magnification, locate the anterior extra lateral nucleus with the vessels as landmarks.

Illuminate it with fluorescent light while searching for a labeled axon near the surface, using a one millimeter outer diameter 0.58 millimeter inner diameter. Bo silicate capillary glass with filament. Pull a recording electrode with a five millimeter long narrow shank and a tip diameter of about one to 2.4 micrometers.

Fill the electrode with filtered hickman's ringer solution. The final tip resistance should be in the range of 16 to 155 mega ohms. Place the electrode in an electrode holder with a pressure port.

Connect it to the head stage mounted on a manipulator. Then connect the head stage to an amplifier. An analog to digital acquisition device not shown connects the amplifier to the computer.

After that, apply 30 millibar positive pressure to the electrode line. Advance the electrode slowly towards the axon from the tissue surface. As the electrode is close to the axon, the positive pressure should cause a slight but noticeable movement of the axon while the electrode is next to the axon.

Record the activity during test stimuli presentation. Although an electrical artifact confirms proper recording and stimulation, there should be no action potentials. Release the outward pressure in the electrode and repeat the stimulation and recording.

Then apply slight suction causing the axon to move into the electrode. Repeat the stimulation and recording again. Now you should see some action potentials in response to the stimulations.

Once action potentials are visible, close the pressure line. When all the desired recordings are complete, switch the tank water back to the respiration solution with MS 2 22 at a hundred milligrams per liter to ensure the fish is completely anesthetized. No EODC should be detected for at least 10 minutes before euthanizing the fish.

Here are five sample traces showing the action potentials evoked by a 0.1 millisecond six millivolt per centimeter, monophasic contralateral positive transverse square pulse stimulus, and here is a raster plot showing the spike times during 20 repetitions of a 75 millisecond recording window for the same unit stimulated at times zero with six millivolts per centimeter stimuli. The range of durations is listed on the right. Shown here is a stimulus duration tuning curve that quantifies the responses as spikes per stimulus repetition.

Once mastered, the surgery and dye injection shown in this technique can be done in one hour if it is performed properly, and recordings can be initiated two to three hours after dye injection. While attempting this procedure, it's important to remember to carefully monitor the health of the experimental animal Following this procedure. Other methods like two photon imaging can be performed in order to record from neurons in even deeper brain structures.

After watching this video, you should have a good understanding of how to label and record single unit responses from an identified population of neurons.