A Time Differential Staining Technique Coupled with Full Bilateral Gill Denervation to Study Ionocytes in Fish

The overall goal of this procedure is to investigate the proliferation of brachial cytes in the gills of goldfish, lacking extrinsic innervation. This is accomplished by first placing the fish in a mitochondrion specific dye bath to stain the preexisting mitochondria, rich brachial cytes. The second step is to perform full bilateral denervation of the gills.

Next, the fish are euthanized. The gills are harvested. An cytes are labeled using an antibody against sodium potassium at TPAs.

Then the final step is to image and count preexisting and newly differentiated cytes. Ultimately, this time differential staining technique with bilateral guild denervation is used to show the proliferation and distribution of gill cytes in goldfish without any nervous input from the central nervous system. So the main advantage of this technique is that it can be used to understand the dynamic uptake of ions and examine the redistribution of cytes in the GI epithelia of fish.

First, prepare one millimolar mitral tracker red stock solution by dissolving 50 micrograms in 94 microliters of dimethyl sulfoxide, and protect from light. Next, prepare three to six dark boxes with a maximum volume of 600 milliliters. Fill each box with 400 milliliters of system water and place an airstone in each to provide a source of oxygen.

Place goldfish weighing 30 to 40 grams into the boxes. After 30 minutes, add the Mitre tracker solution to yield a full concentration of 0.1 micromolar dye and 0.01%DMSO. Bathe the fish for four hours.

After four hours, turn on the water flow to the boxes and allow the dye to flush out. If these are control animals, recover the fish for three to five days, then proceed to immunohistochemistry, otherwise proceed to the full bilateral denervation procedure. First, anesthetize the fish by placing the fish in an aerated benzocaine water bath.

Once breathing has ceased, place the fish on a surgery table and insert a tube into the buccal cavity To irrigate the gills with aerated anesthetic solution, position the fish so that the head is tilted slightly downward to allow for better access to the area behind the fourth gill arch. Gently lift the operculom with a straight standard pattern forceps, and then place tissue retractors between the operculom and the inside of the head. Carefully open the tissue retractors to keep the operculom away from the head and keep the gills exposed.

Rest the retractor handles next to the head To provide access to all four gill arches. Check that the anesthetic solution is irrigating the gills and place curved or straight standard pattern forceps between the fourth gill arch and the back of the head. Gently open the forceps to create tension in the ligament.

Attaching the fourth gill arch to the head with a pair of number five forceps. Create a two to three millimeter opening by piercing the epithelium, connecting the dorsal end of the gill arches to the auricular cavity. Be careful not to go in too deep because there is the risk of damaging a major blood vessel.

Next, hold a small cotton bud with number five forceps, and use it to carefully expand the incision to expose the ninth and 10th nerves. Free the nerves from any connective tissue by using the number five forceps. Again, taking care not to damage blood vessels.

Once the nerves have been exposed and identified, use the curved spring scissors to carefully cut the nerves while holding the incision. Open with the straight standard shape forceps. After severing the nerves, gently retract the curved forceps and remove the tissue.

Retractor sutures are not required because the incision should close on its own Within 24 to 48 hours. After repeating the procedure on the other side of the head, switch the irrigation of the gills from anesthetic to fresh aerated water and allow the fish to recover from anesthesia. Once auricular movements have resumed, move the fish into experimental tanks to recuperate for at least 24 hours.

Finally, perform a sham procedure on a separate set of fish by piercing the epithelium behind the fourth gill arch without severing the nerves before euthanizing the fish. Place three to four milliliters of 4%PFA into a scintillation vial. Use one vial per gill arch.

In addition, take a small whey boat and fill it with one XPBS. This will be used to wash the tissue after it has been excised. Keep all solutions on ice.

After euthanizing the fish with an overdose of benzocaine, use blunt forceps to lift the perm on one side of the head and curved scissors to sever each end of the brachial gill basket. Next, use blunt forceps to carefully pick up the gill by the Rackers and lift it out of the auricular cavity. Immediately wash the gill in ice cold one XPBS to remove excess benzene and blood and place the excised gill into a vial filled with 4%PFA fix all excised gills overnight at four degrees Celsius after fixation.

Wash off X-S-P-F-A in one XPBS, and place the tissue in a two milliliter bullet tube filled with 1.5 milliliters of 1%Tritton x. Place the bullet tubes on a shaker for six hours at room temperature or overnight at four degrees Celsius. To perme the tissue, prepare a one to 250 dilution of each primary antibody by adding four microliters of both the NKA and ZN 12 primary antibodies into 992 microliters of one XPBS.

Make sure that both primary antibodies have been raised in the same host. Aspirate the Triton X solution and without washing the tissue first and the primary antibody solution and incubate on a shaker for six hours at room temperature or overnight at four degrees Celsius. Following the incubation, aspirate the primary antibody solution and wash the primary antibody three times for three minutes each.

Using one XPBS. Prepare the secondary antibody at a one to 200 dilution by mixing five microliters of stock secondary antibody into 995 microliters of one XPBS. After aspirating PBS from the last wash, apply the secondary antibody and incubate for six hours at room temperature or overnight at four degrees Celsius on a shaker.

Finally, remove any excess secondary antibody by washing the tissue three times for five minutes each as before, after the washes mount the tissue by first placing a gill in a 200 microliter drop of one XPBS on a flat microscope slide. To ensure that the tissue does not desiccate. Separate the gill hemi Brinks with curved micro scissors.

Then place a drop of one XPBS and a drop of mounting media into the depression of a concave slide. Place the separated hemi branks into the concave slide with the leading edge of the filament facing upward and cover with a cover slip dab the edges of the cover slip with nail polish. In order to prevent the cover slip from moving around and displacing the tissue, allow the tissue to settle to the bottom of the concave slide for 10 to 15 minutes before imaging.

For each gill arch, select six gill filaments at random for imaging producing six images per gill arch. Use conventional confocal microscopy to image the tissue by taking one to three micrometer optical slices in the following images, cytes indicated by arrowheads were stained with the alpha five antibody and nerves indicated by arrows were stained with the ZN 12 antibody. This confocal image of a gill filament from a control fish shows extrinsic innervation with extensive lamella branching of a central nerve bundle, presumably originating from the ninth and 10th nerves as seen on the inset image.

Some of the cytes indicated are also innervated. This image shows a gill filament two days after full bilateral denervation. There was a reduction of the central nerve bundle while the laminar innervation appeared largely intact.

Here, a gill filament shown five days after full bilateral denervation demonstrates that extrinsic innervation is now largely absent. Qualitative analysis suggests that full bilateral denervation causes degradation of the extrinsic gill innervation while maintaining nerves with cell bodies within the gill filament, creating a network of nerves through the filament and into the lamely. This procedure can be coupled with other methods such as measuring ion fluxes, using radioactive ions like sodium 22 in order to answer additional research questions.