This protocol demonstrates a novel procedure for sequential immunofluorescence and immunohistochemistry on cryosections from early stage zebrafish embroys, which enables precise co-localization analyses in specific cell populations. The main advantage of this protocol is it's flexibility. It combines immunofluorescence and immunohistochemistry techniques using a single cryosection to maximize tissue morphology, co-localization of expression and antibody compatibility.

This protocol could be applied to other fish or amphibious models as these researchers often face similar complications with antibody availability and often perform similar types of experiments. Working with early stage embryo cryosections can be tricky since they are small and cryosectioning can be challenging, but advanced preparation and practice will help alleviate any struggles in this method. There are multiple steps in our protocol that require particular handling and care and can be difficult to master without seeing someone demonstrate the techniques.

To begin, transfer previous fixed and prepared forty eight hour post fertilization chimeric zebrafish embryos from a tube with 15 25 OCT mixture to a plastic mold using forceps minimizing any transfer of the mixture. Fill the mold approximately half with OCT medium and gently mix the embryos in it. Prepare labeled plastic molds and transfer desired embryos into the empty labeled plastic molds, minimizing carryover of OCT medium.

Then gently fill with OCT medium to the top of the molds with embryos. Working under a light stereo microscope for visualization, use needles to arrange embryos in the desired orientation. Then place the prepared molds in an insulated container with a metal platform and freeze them on dry ice.

Place an ice bucket gently over the metal platform to create a cold chamber and freeze for approximately 30 minutes. Using a cryostat set at minus 20 degrees Celsius, cut the embedded embryos in 10 to 12 micrometer thick cryosections while checking the depth of the section periodically on a microscope to monitor location and tissue. Place the sections on charged glass slides and air dry them at room temperature overnight.

Wash the slides three times for five minutes in 1X PBS in an appropriate container. Lay the slides on a flat surface in a humid chamber. Use a barrier pen to outline the sections to keep liquid on the slides.

Pipette 200 microliters of block buffer per section and incubate in block buffer for two hours at room temperature. Prepare the primary antibody dilution and block buffer and mix well by pipetting. Gently tip the slides to drain off the block buffer and return to the humid chamber.

Pipette 200 microliters of primary antibody solution per section onto the slides and 200 microliters of block buffer onto to the appropriate sections as control. Incubate the slides at four degrees Celsius overnight in a humid chamber filled with deionized water making sure to seal the edges of the chamber to help retain moisture. Wash the slides three times for five minutes in 1X PBS and in an appropriate container.

Prepare the secondary antibody dilution during washing and block buffer and mix well by pipetting. After laying the slides on a flat surface in a humid chamber, add 200 microliters of secondary antibody solution per section. Incubate the slides in secondary antibody solution at room temperature in the dark for 30 minutes.

Wash the slides three times for five minutes in 1X PBS in an appropriate container. After placing the slides on a flat surface, add the nuclear staining solution onto each section and incubate covered for 10 minutes. After draining off the nuclear staining solution, mount the slides with non-hardening fluorescent mounting media and a glass cover slip.

Making sure to keep them in the dark at four degrees Celsius until imaging. Place the slides in individual containers with 1X PBS and store flat at four degrees Celsius overnight, while gently agitating to loosen the cover slips enough so they will come off when agitated. Make sure not to press down on the cover slip or attempt to remove the cover slip manually but wait until they come off with minimal forced movement.

After the cover slips have been removed, gently transfer the slides into a container with fresh 1X PBS. Remove the 1X PBS and incubate the slides in 1X tris-buffered saline with 0.1%of a non-ionic surfactant three times for five minutes at room temperature. Incubate the slides in 3%hydrogen peroxide solution at room temperature for 15 minutes.

Then lay the slide flat and add block buffer drop wise to the surface. Gently tip the slides to drain off the block buffer. Dry the area around the sections and place the slides flat in a humid chamber.

Add 200 microliters of primary antibody solution per section onto the slides and incubate in humid chamber at four degrees Celsius overnight. Gently tip the slides to drain off the primary antibody solution and wash two times for five minutes in 1X TBST. Then apply ready to use background reducing blocking reagent drop wise to each section, and incubate in humid chamber at room temperature for 20 minutes.

Gently tip the slides to drain off the block buffer. Carefully dry the area around the sections and place the slides flat in a humid chamber. Apply a ready to use secondary antibody solution to each section drop wise and incubate in humid chamber at room temperature for 30 minutes.

Gently tip the slides to drain off the secondary antibody solution and wash two times for five minutes in 1X TBST. After drying the area around the sections, place the slides on a flat surface and add 200 microliters of the HRP chromogenic substrate to each section. Start a timer when the substrate has been applied to the first slide and incubate at room temperature for three minutes.

Drain off the substrate solution. Rinse the slides briefly in an appropriate container with 1X PBS and wash them in deionized water twice for five minutes with gentle agitation. Counterstain the slides by placing them in hematoxylin staining solution for up to 30 seconds and wash three times for five minutes each in deionized water.

Incubate the slides in a container containing Scott's Tapwater for one minute and then wash again three times for five minutes in deionized water. Dehydrate the slides through a series of ethanol grades diluted in deionized water and xylene in a chemical hood. After removing the slides from xylene immediately add toluene based mounting medium and place a cover slip.

To ensure success while cover slipping, it is important to work from one side of the slide to the other while slowly lowering the cover slip to prevent bubbles which would obscure the tissue. Finally, visualize and image the slides using a compound light microscope and digital camera at 100X magnification. Specific antibodies used here to simultaneously detect proliferating cells and donor cells identified and quantified donor cells that are actively proliferating.

After both immunofluorescence and immunohistochemistry it is essential to generate high quality images in order to accurately identify individual cells. An image analysis program was used to perform image overlay. When attempting this procedure it is most important to adequately stain and counter stain slides during immunohistochemistry.

After this procedure additional methods can be performed as modifications to the original protocol, such as using different antibody combinations, tissue types, or species. Images can also be analyzed in a variety of ways to obtain relevant data. This technique allowed us to look at co-localization and multiple genetic backgrounds as a way of investigating cell to cell interactions and could be applied similarly to other techniques requiring detailed co-localization analysis.

Many of the reagents in the immunohistochemistry are hazardous and should be treated accordingly. Work with ethanol, xylene, and mounting stiff should be performed in the hood. DAB waste should be disposed of as hazardous waste and post DAB washes should be bleached.

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Antibody Compatibility