Determining Viral RNA Distribution in Host Cells by Fluorescence In Situ Hybridization

Begin with a slide containing permeabilized, virus-infected host cells, where viral RNAs are distributed between the nuclear and cytoplasmic compartments.

Add a pre-hybridization buffer and incubate under humid conditions.

In the buffer, blocking reagents block nonspecific binding sites, while RNase inhibitors inactivate RNases and preserve RNA integrity.

Remove the buffer and add denatured, digoxigenin-tagged oligonucleotide probes complementary to viral RNA.

Incubate in the dark to allow probe-RNA hybridization, then wash repeatedly.

Add a fixative to stabilize the probe-RNA hybrids, then wash repeatedly. Add a detergent-supplemented buffer to maintain cell permeability.

Add a green-fluorophore-conjugated anti-digoxigenin antibody that interacts with digoxigenin, then wash repeatedly.

Fix the cells to preserve the complexes, then wash again.

Add a DNA-binding dye to stain the nuclei. Wash excess dye repeatedly.

Mount with a coverslip and seal it.

Under a confocal microscope, visualize the green fluorescence, reflecting the distribution of viral RNA between the host cell nucleus and cytoplasm.

After three PBS washes as demonstrated, fix the samples again in 4%formaldehyde in PBS for 10-15 minutes before a second permeabilization as demonstrated. Then cover the slide with aluminum foil to preserve the fluorescent signal and to prevent photobleaching.

And wash the cells with 2x saline sodium citrate, before applying 45 microliters of hybridization solution. After one hour at 37 degrees Celsius in a humidified chamber, add distilled water to the antisense oligonucleotides of interest to bring the denaturation volume to 10 microliters. Denature the oligonucleotides at 95 degrees Celsius for five minutes, before adding 35 microliters of fresh hybridization solution per slide.

Then incubate the samples from 10-24 hours in the humidified chamber at 37 degrees Celsius, protected with foil. The next day, wash the cells with two, 10 minute. 2X saline sodium citrate washes at room temperature, followed by one wash in 1X saline sodium citrate for 10 minutes, at 25 degrees Celsius.

After the last wash, fix the cells with pre-chilled 4%formaldehyde in PBS, for 10-15 minutes on ice, followed by three washes with fresh PBS. Next, permeabilize the samples as demonstrated, followed by incubation with anti-digoxigenin-FITC and pre-chilled 0.1%BSA in PBS, for one hour at four degrees Celsius. At the end of the incubation, wash the slides three times in fresh PBS, and fix the samples with pre-chilled 4%paraformaldehyde in PBS, for 10-15 minutes at four degrees Celsius.

After three washes in PBS, label the nuclei with 0.4 micrograms per milliliter of DAPI and pre-chilled 0.5%Triton-X in PBS for 15 minutes on ice, followed by three final washes in PBS. Mount slides with fluorescent beads, as experimentally relevant and in appropriate mounting medium. After removing excess mounting medium with sterile wipes, seal one coverslip to each slide with multiple coats of clear nail polish, and use a fluorescence microscope to confirm successful execution of the experiment.

Images may then be taken on a confocal microscope to collect images of the samples within an hour to a week of mounting, at a 630 times magnification.