This article details a protocol for staining extracellular DNA in tissue samples. The method involves several steps including deparaffinization, antigen retrieval, and antibody staining to visualize DNA using fluorescence microscopy.
A dying cell releases a fraction of DNA outside the cell called the extracellular DNA. To stain extracellular DNA, begin by taking a formalin-fixed paraffin-embedded, or FFPE, tissue section on a glass slide. Heat the glass slide for the desired duration. This facilitates the attachment of tissue on the slide surface and softens the surrounding paraffin.
Immerse the slide in deparaffinization liquid to completely dissolve the paraffin layer around the tissue. Next, dip the slide in decreasing concentrations of ethanol to rehydrate the deparaffinized tissue. Following rehydration, submerge the slide in boiling antigen retrieval solution. This treatment unmasks the epitopes that were altered during the fixation process.
Overlay the slide with primary antibodies and allow them to bind to specific unmasked epitopes present on the cell. Then, add fluorescently-tagged secondary antibodies, which bind to the primary antibodies. Dip the slide in Sudan Black solution to reduce background autofluorescence.
Finally, place a drop of mounting medium supplemented with DAPI over the tissue section and seal it with a coverslip. Incubate to allow the DAPI molecules to stain the DNA. Visualize the prepared tissue section using a fluorescence microscope. Fluorescently labeled epitopes in the cell help in differentiating the nuclear DNA present inside the cell from the extracellular DNA.
For extracellular DNA staining, first, heat the samples of interest in the slide rack in 60 degrees Celsius for 60 minutes before immersing the slides in two 40 minute changes of solvent in a fume hood. After the second immersion, rehydrate the samples two times in 100% ethanol and one time in 70% ethanol for 5 minutes per treatment.
After the last treatment, use tongs to place the slides into boiling antigen retrieval solution in a pressure cooker on high for 10 minutes. After unmasking the antigen epitopes, transfer the pressure cooker from heat into a sink and immediately run cold tap water over the lid.
Allow the slides to equilibrate for 20 minutes in the antigen retrieval solution before washing the samples two times in 0.01 molar PBS on an orbital shaker for 5 minutes per wash. After the second wash, use a hydrophobic pen to draw circles around the kidney tissue samples and block any non-specific staining with 60 microliters of 10% chicken serum in 5% bovine serum albumin for 30 minutes at room temperature.
At the end of the incubation, carefully replace the blocking solution with 60 microliters of the primary antibody of interest in a humidity chamber overnight at 4 degrees Celsius. The next morning, wash the slides on an orbital shaker in PBS for 2 minutes per wash before adding 60 microliters of an appropriate fluorophore-conjugated secondary antibody to each slide for a 40-minute incubation at room temperature.
At the end of the incubation, wash the slides in PBS as demonstrated, followed by treatment in 0.3% Sudan Black in 70% ethanol to quench any potential formalin-induced autofluorescence. After 30 minutes, wash the slides in tap water to remove any precipitate and immerse the slides in PBS for 10 minutes to prevent any further Sudan Black precipitate formation.
At the end of the incubation, mount the slides onto confocal glass coverslips with 360-microliter drops of mounting solution supplemented with DAPI and seal the coverslips with nail polish. Allow the slides to cure for 24 hours at room temperature. Store them at 4 degrees Celsius protected from light until imaging by confocal laser scanning microscopy according to standard protocols.
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