This article details a protocol for visualizing DNA damage response proteins at double-strand break sites using immunofluorescence microscopy. The method involves fixing and permeabilizing human mononuclear cells, followed by antibody incubation and fluorescence imaging.
To visualize specific DNA damage response proteins at DNA double-strand break, DSB, sites using immunofluorescence microscopy, begin with microscope slides containing human mononuclear cell monolayers with DSBs. Fix the cells with paraformaldehyde.
Treat with a non-ionic detergent, permeabilizing membranes for accessing intracellular targets. Add a protein-containing blocking solution, blocking cells' non-specific binding sites. Incubate with primary antibodies that bind to the target response proteins.
Treat with a secondary antibody mixture tagged with different fluorophores to bind specifically to their respective primary antibodies attached to the proteins. Stain DNA with DAPI. Add mounting medium; mount the coverslip.
During fluorescence microscopy, the incident light beam reaches the excitation filter allowing light of a specific excitation wavelength to pass through. Upon reaching the dichroic mirror — a beam splitter — the excitation light selectively reflects towards the objective lens, which focuses the light onto the sample.
The incident excitation light excites the fluorophore's electrons to a higher energy state. On returning to the ground state, the electrons emit light at a longer wavelength, collected by the objective lens, and passed back to the dichroic mirror.
The mirror allows the emitted fluorescence light to pass through the emission filter, which allows only the desired fluorescence light to reach the detector. Process the images. Analyze the different colored fluorescent foci to visualize the co-localization of the DNA damage response proteins at the DSB site.
Prepare two cytospins with mononuclear cells of the patient samples by centrifugation of 1 x 105 cells for each preparation. Fix the cells with 200 microliters of 4% PFA for 10 minutes.
After fixation, wash the cells gently three times with 30 milliliters of PBS for five minutes each on a shaker. Then, permeabilize the cells with 200 microliters of 0.1% Octoxinol-9 for 10 minutes. Wash the cells gently three times with 30 milliliters of 5% blocking solution for five minutes each on a lab shaker. Block the cells in 30 milliliters of fresh 5% blocking solution for one hour.
The fixation and permeabilization of the cells with 4% paraformaldehyde and 0.1% Octoxinol-9 preserved the gamma H2AX and 53BP1 foci distinctly.
Incubate one preparation of the cells with a mouse monoclonal anti-gamma H2AX antibody and the other preparation of the cells with a mouse monoclonal anti-gamma H2AX antibody and a polyclonal rabbit anti-53BP1 antibody overnight at 4 degrees Celsius.
The use of proven anti-gamma H2AX and anti-53BP1 antibodies is highly recommended.
After incubation, wash the cells gently three times with 30 milliliters of 2% blocking solution for each five minutes on a lab shaker.
Next, incubate the first preparation of the cells with an Alexa488-conjugated goat anti-mouse secondary antibody diluted 1 in 500 in 2% blocking solution. Incubate the second preparation of the cells with an Alexa488-conjugated goat anti-mouse secondary antibody and an Alexa555-conjugated donkey anti-rabbit secondary antibody diluted 1 in 500 in 2% blocking solution.
Wash the cells gently three times with 30 milliliters of PBS on a laboratory shaker. Remove the PBS and mount the cells with mounting medium. Cautiously, put a coverslip on top of the mounting medium, so that no air bubbles are embedded. Wait at least three hours for the mounting medium to harden before analyzing the cells by fluorescence microscopy.
Finally, analyze the gamma H2AX and 53BP1 foci in the cell nuclei with a fluorescence microscope equipped with filters for DAPI, Alexa488, and Cy3 during imaging at a 100x objective magnification.
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