The overall goal of this experiment is to evaluate the patho-histology of the zebrafish larva liver caused by acute ethanol insults. This method can help address key questions in the field of alcoholic liver disease such as the histological feature of acute alcohol liver injury and to what extent the responses of zebrafish liver to acute alcohol injury resembles those seen in humans. The main advantage of this technique is that our ethanol treatment induces more robust responses than other published methods.
Also, our histology protocol improves tissue integrity on the sections. Begin by selecting up to 40 96-hour post-fertilization larvae with inflated swim bladders using a Pasteur pipette. Split them evenly into two new Petri dishes.
Remove as much residual egg water as possible, then add egg water containing 2%ethanol to one dish at a density of one larva per milliliter. Add the same amount of egg water without ethanol to the other dish to serve as the control. Keep the control and ethanol-treated larvae in the fish room for 24 hours.
After the 24-hour period has elapsed, collect the larvae in a 1.5-milliliter centrifuge tube with no more than 20 larvae per tube. While working in the chemical hood, remove as much liquid as possible from the larvae, and then add at least one milliliter of Dietrich's fixative. Allow the fish to fix on a nutator at room temperature for at least 24 hours.
Set out the tissue cassettes and two blue biopsy pads per cassette. Place one biopsy pad at the bottom of each cassette. Label each cassette in pencil clearly identifying the sample.
While working in the fume hood, remove the fixative from the tubes, and wash the larvae three times for five minutes each time with one milliliter of PBS. Place the tubes on a nutator at room temperature while washing. During the washes, heat 3%agarose in water using a microwave to melt the solid.
Heat for 30 seconds at a time, and watch carefully to minimize boiling. Once dissolved, place the liquid agarose on a hot plate set to 90 degrees Celsius with gentle stirring. Using a transfer pipette, transfer up to eight larvae to a plastic histology mold, and remove as much PBS as possible.
Using a transfer pipette with the tip cut off, completely fill the mold with 3%agarose, then use an insulin syringe to gather the larvae to the center of the mold, and push them to the bottom. For sagittal sections, position the larvae in a line with the heads toward the top of the mold. To ensure that the livers are oriented in a consistent manner, turn the larvae so that the left side of the body is facing down, and flat against the bottom of the mold.
After allowing the agarose to set for four to five minutes, remove the agarose block from the mold, and use a razor blade to trim the agarose around the larvae. Stand the block on end, and cut the thickness in half so that the final block is roughly two to three millimeters thick. Transfer the small agarose block to the prepared tissue cassette, and place the second biopsy pad on top of the block.
Close the cassette, and place the fully assembled cassette into a sealable container with freshly prepared 70%ethanol in double-distilled water. After paraffin embedding the tissue, use a microtome to remove excess paraffin covering the tissue by sectioning five microns at a time until the tissue is exposed at the surface of the paraffin block, then stop and discard all the sections that are cut. Soak the blocks face down in PBS, and place it four degrees Celsius overnight.
Then next day, remove one block at a time from the PBS, and cut five micron sections using a microtome. Separate the ribbon of sections from the blade by gently pulling the last section away from the blade using forceps or a brush. Use the forceps to pick up the ribbon by the last section, and transfer it to a water bath at 42 degrees Celsius.
Allow the ribbons to float on the surface of the water for at least five minutes. Use the forceps to tease apart sections into smaller groups to fit onto the slides. Put a charged slide into the water at a 45-degree angle, and carefully position it underneath the group of sections to be collected.
Carefully lift the slide from the water, and allow the sections to attach to the slide. Blot any excess water from the sections using lint-free tissue. Place the slide in a slide holder or box.
Continue to section the block until the desired tissue has been collected. When finished, bake the slides in a 55 degrees Celsius incubator or oven for three to 16 hours to melt the paraffin. After the allotted time, check that the paraffin has melted, and allow the slides to cool before staining.
Deparaffinize the slides by immersing them in 100%xylene twice for 15 minutes each time, then rehydrate the slides through an ethanol gradient. For each solution, dip eight to 10 times for two seconds per dip until the liquid runs cleanly off the slides. Place the slides in 100%filtered Harris Hematoxylin for four minutes.
After four minutes, quickly transfer them back to the container of deionized water. Run deionized water into the back corner of the container farthest away from the sections. Empty the container periodically until the water is no longer purple.
Quickly check the Hematoxylin intensity on a dissecting microscope using gooseneck lights. Return the slides to 100%Hematoxylin for one minute if further staining is required. Once the desired Hematoxylin staining has been obtained, dip the slides twice in 0.05%hydrochloric acid, then immediately transfer them back to the container with clean deionized water.
Empty and refill the container with water twice. Transfer the slides to two containers of 95%ethanol for 30 seconds each. Place the slides into the Eosin Y-Phloxine B solution for two minutes.
After the two minutes, transfer the slides back to the previous 95%ethanol container, then check the intensity of the color under the dissecting microscope. If the staining is not sufficient, return to the Eosin solution for 30 seconds. Repeat as necessary.
When staining of the desired intensity has been obtained, transfer the slides to 100%isopropanol for 15 seconds. Replace with fresh 100%isopropanol, and place the slides back in the isopropanol for another 15 seconds. Repeat this process for a total of six isopropanol washes.
After immersing in 100%xylene for three minutes, remove one slide. Add sufficient mounting medium to cover the sections, and then dip the slide in 100%xylene. Apply a cover slip to the slide, and blot any excess mounting medium on a paper towel until only a thin line is seen, then dip a tissue into the xylene, and wipe the back of the slide to remove any medium that has dripped.
Place the slide flat on a sturdy but mobile surface like a piece of cardboard, and allow the xylene to evaporate in the hood for 10 minutes. Leave the slides at room temperature overnight to allow the mounting medium to harden before imaging. This 120 hours post-fertilization zebrafish larvae liver was immersed in Dietrich's fixative at room temperature for 24 hours.
Dietrich's fixative provides the optimal fixation of this tissue. With 10%formalin fixation, the hepatocytes often lose their cytoplasm. The tissue is not stained properly with Eosin, and appears purple.
After fixation with 4%PFA, gaps are seen between the hepatocytes. 4%PFA causes the liver tissue to shrink so there appears to be a large gap between the liver and the surrounding tissues. The dashed line marks the border of the surrounding tissues.
This higher power image shows H&E staining of the liver in a wild-type untreated control larva that was part of an ethanol treatment study. Again, this animal was fixed with Dietrich's fixative at 120 hours post-fertilization. This image shows the effect of treatment with 2%ethanol from 96 to 120 hours post-fertilization on the liver.
This acute ethanol treatment causes hepatic steatosis, and blood vessel swelling. The arrows point to the hepatocytes with excessive deposition of round droplets, and the asterisks mark the swollen hepatic blood vessels. While attempting this procedure, it's important to remember to perform the ethanol treatment in a fish facility with a light-dark cycle.
For H&E, use Dietrich's fixative for an optimal tissue preservation. Also remember to check the color intensity frequently to avoid over-staining or under-staining.
