An Immunolabeling Technique to Visualize the Subcellular Localization of Proteins

Begin by washing a fixed heart section in a sodium cacodylate buffer that stabilizes the tissue structure. Immerse the section in osmium tetraoxide to stain the membrane lipids.

Treat with sodium acetate to maintain an optimum pH for staining.

Add uranyl acetate to stain biomolecules, providing better contrast during microscopy.

Dehydrate the tissue in increasing concentrations of ethanol followed by acetone.

Embed the tissue in a resin, obtain ultrathin sections, and transfer them onto a microscopy grid.

Introduce sodium meta-periodate to remove the osmium tetraoxide, unmasking the cellular protein. 

The compound also oxidizes glycoprotein, creating reactive aldehyde groups, which are neutralized by glycine treatment.

Introduce a blocking buffer, preventing non-specific antibody binding.

Add primary antibodies specific to the target protein followed by biotinylated secondary antibodies.

Add streptavidin-conjugated quantum dots that bind to biotin.

Under an electron microscope, quantum dot-labeled regions exhibit higher contrast, aiding precise subcellular localization of the target protein.

After 24 hours of fixation, wash the tissue in 0.1 molar sodium cacodylate buffer twice for 20 minutes. Remove the tissue from sodium cacodylate buffer and immerse in 2% osmium tetroxide solution for 4 hours at room temperature.

After osmication, immerse the tissue in 2% sodium acetate solution for 10 minutes at room temperature. Next, immerse the tissue in 2% uranyl acetate solution for 1 hour at room temperature. After uranyl acetate staining, dehydrate the tissue sequentially through the graded alcohols and acetone.

Embed the dehydrated tissue in low-viscosity epoxy resin as described in the manuscript. Put the tissue into fresh resin in 8-millimeter micro-molds, molds, and cure the embedded tissue at 70 degrees Celsius overnight. After finding the area of interest using an ultra 45-degree knife, produce pale gold ultra-thin sections. Place these ultra-thin sections on the dull side of a 200-mesh copper grid.

Start the staining by unmasking the antigen by putting 20 microliters of etching solution sodium metaperiodate on a clean paraffin film. Place the dried grid with tissue sections on the droplet of the etching solution. Leave the section grid on the solution for 30 minutes at room temperature. Wash the etched tissue sections by placing them on a droplet of distilled water for 60 seconds.

Block the residual aldehydes by placing the section grid on a droplet of 0.05 molar glycine solution for 10 minutes at room temperature. Blot the grid's edges on filter paper to remove the residual glycine solution. Place the section grid in 10 to 20 microliters of blocking solution for 25 minutes at room temperature.

Blot the grid edges on filter paper, and place the grid sections on antibody diluent for conditioning at room temperature for 10 minutes. Incubate the grid sections with primary antibody for 1 hour 30 minutes in a humidified chamber. Blot-dry the grid, and wash the grid sections with antibody diluent twice for 5 minutes each.

Incubate the grid sections with biotinylated secondary antibody for 1 hour in a humidified chamber. Once the grid is blot-dried, wash the grid sections with antibody diluent twice for 5 minutes each. Incubate the grid sections in commercially available streptavidin-conjugated QD for 1 hour in a humidified chamber at room temperature. Prevent exposure to light by covering the chamber with aluminum foil. After blot-drying the grid edges using filter paper, wash the grid sections by placing them on water droplets for two minutes, and blot the grid edges to dry.