膜片钳记录在内耳毛细胞的分离斑马鱼

The aim of this procedure is to demonstrate the methods for obtaining healthy, intact hair cells from the auditory and vestibular end organs of adult zebra fish so that they can be used in patch clamps. Studies aimed at characterizing the biophysical properties of their voltage gated channels. This is accomplished by first removing the brain from the fish and then taking out the ventral portion of the brain case that contains the labyrinths.

The second step is to break apart the bowl of bone in order to isolate the labyrinths that contain the end organs, the S lega, utricle, and semicircular canals. Next, a dog hair is used to lift off from the end organs, that sheets of epithelia that contain the hair cells. The final step is to isolate individual hair cells by tri curating the epithelial sheets with two dog hairs.

Ultimately, patch lamp recordings can be made on isolated hair cells to characterize the properties of voltage gated channels. The main advantage of this technique over existing methods is that it shows how to obtain viable hair cells from the genetically tractable zebrafish model organism. This method can be used to help answer key questions in the field of auditory and vestibular physiology, such as those that seek to understand the mechanisms that control the release of neurotransmitters from cells that mediate hearing and balance.

Begin this procedure by preparing six different solutions according to the accompanying manuscript. Next label and fill 4 35 millimeter Petri dishes approximately halfway with the solutions indicated in the accompanying manuscript. After that, prepare a dissecting dish by cutting a fish shaped cavity in a 60 millimeter Petri dish filled with cured sil guard.

This will prevent the animal from tipping over during the dissection. Prepare at least two cell isolation tools by gluing the follicle end of a dog hair to the end of a glass pasta pipette with super glue, allowing the hair to extend beyond the end of the pipette by approximately 0.5 centimeters. Now, sacrifice one adult zebra fish by immersing it in a beaker containing normal zebra fish ringers or NZR and trica.

Visually monitor the gills until all the auricular movements have ceased. Wait at least an additional 10 minutes before removing the fish and rinsing with fresh NZR.Afterward. Place the fish in the dissecting dish with its dorsal side up.

Insert one standard sewing straight pin through each eye socket, and a third pin through the dorsal ventral axis. About one third to one half of the distance from head to tail under a zoom stereo dissecting microscope, expose the brain and a short segment of the spinal cord by removing the skull roof from a point approximately 0.5 centimeters cordal to the level of the gills to the nose of the fish using a pair of spring scissors. Then cut the spinal cord and lift the brain with the fine forceps while cutting the nerves and other attachments.

After that, remove and discard the brain. The remaining ventral portion of the skull capsule forms a bowl that contains the inner ear organs. Observe the prominent white opaque OTA otoliths in the legina and sculli located symmetrically at the quarter end of each inner ear and the laterally positioned utricle otoliths located.

More tally. These structures are useful landmarks to help in the intact removal of the labyrinth with a pair of fine forceps and spring scissors. Remove the bowl of bone by cutting all the ventral and lateral attachments while gently lifting it outta the head.

Place it in the Petri dish containing low calcium solution or low cas next. Crack apart the bowl at its midline, and separate the right and left halves by inserting the points of fine forceps into the center and prying the halves apart. Then remove the two labyrinths from the bone.

Inspect the labyrinths and identify the auditory and vestibular end organs. The semicircular canals, uracil, sculli, and legina. Carefully remove the odor lifts from the legina and utricle using fine forceps.

In this procedure, transfer the end organs to the dish containing locas and propane. Using a plastic transfer pipette incubate these structures for 30 minutes at room temperature. At the end of the incubation period, transfer the structures to the dish containing NZR and BSA and incubate for at least 30 minutes.

More to prepare for cell isolation. Transfer one of the end organs, for example, the legina to the dish containing NZR. This dish is then placed on the stage of a stereo zoom dissecting microscope.

Use a dog hair to gently lift off the macular the sheet of pinkish epithelial tissue that contains the hair cells. The mace can also be removed from the sculli and uracil, and the kae can be isolated from their locations inside the semicircular canals. Using two of the cell isolation tools prepared earlier, Tate the macular to generate a debris field that will contain the hair cells.

Allow the cells at least five minutes to settle onto the bottom before moving the dish. Next, transfer the dish to the stage of an inverted microscope equipped with face contrast optics. Observe the cells under the 40 times magnification.

Note the diversity in cell morphology. Some cells are avocado shaped while others are long and thin. The presence of apical hair bundles indicates hair cells and phase brightness assures their health.

For perforated patch recordings, prepare an amphotericin containing solution by first placing five milligrams of amphotericin B into a 1.5 milliliter centrifuge tube. Then add 100 microliters of DMSO and immediately vortex the tube for 10 to 20 seconds until all of the amphotericin is dissolved. Next pipette 625 microliters of potassium ion internal solution into a second micro centrifuge tube.

Then add 10 microliters of the amphotericin solution to the potassium ion internal solution and vortex it for another 10 to 20 seconds. Fabricate patch pipettes from filament containing boro silicate glass capillary tubing. Using a multi-stage puller, the pipette should have tipped diameters of approximately two microns and resistances between one and three mega ohms.

Blunt bullet shaped pipettes are preferred as they offer the least access resistance during electrophysiological recordings. After that, use a 28 gauge microfill syringe needle to fill a patch pipette halfway with the amphotericin containing potassium ion internal solution. Affix this pipette to the head stage of a patch clamp amplifier to enable patch clamp recordings.

First, apply positive pressure to the electrode as it is lowered through the air solution interface and down to the bottom of the recording dish near the cells. Next, position the electrode orthogonally to a healthy looking cell. When the electrode is close enough that the cell begins to move away from the outflowing solution, quickly reverse the pressure creating a slight vacuum in the electrode until the cell jumps onto it.

Immediately cease the suction on the electrode to allow the formation of a giga ohm seal with the cell. Apply repetitive hyperpolarizing voltage steps of 10 millivolts to the electrode and observe the capacitive current transience. Continue to monitor the amphotericin induced membrane perforations as the magnitude of the transient grow to their maximum size.

To confirm the establishment of the perforated patch configuration in a healthy cell, elicit voltage gated currents by applying hyperpolarizing and depolarizing steps of potential. Most hair cells have prominent outward currents that rapidly inactivate during maintained depolarization. This figure shows the ventral portion of the brain case after the removal of the brain.

The otoliths associated with the tulo, genus and sculli, as well as those associated with the two utricle can be seen through the thin bone that overlies them after removal of the bowl of bone. The presence of the otoliths helps to identify the location of the labyrinths. The dash line in the cartoon indicates the approximate perimeter of the left labyrinth.

When the labyrinth is isolated, the auditory and vestibular end organs can be identified such as the s, legina, utricle, and semicircular canals. The cartoon illustrates the location of these structures, A, B, and C of the semicircular canals. D is the utricle, E is the s, and f is the lega.

The diversity in hair cell morphology is apparent in this figure of cells isolated from the lega. Cells roughly fall into two classes, either thin or avocado shaped. Healthy cells are phased bright with sharp outlines and obvious hair bundles by contrast.

Dead cells shown here in inset B have a granular black appearance. This figure shows the averaged responses in a cell to three presentations of voltage steps applied in 10 millivolt increments from minus 140 millivolts to plus 70 millivolts with a holding potential of minus 70 millivolts. Note that the inactivating current appears at more depolarized potentials.

Once mastered, electrophysiological recordings can be taken as soon as 90 minutes from the start of the procedure. Furthermore, if the tri step is postponed and the end organs are kept in BS, a solution, healthy cells can be obtained up to four hours after the start of the dissection. This preparation when using combination with widely available zebrafish genetic mutations provides a unique model system to investigate the molecular factors that underlie the mechanisms of hearing imbalance.