Immunostaining to Visualize Viral Antigen and Host Protein Colocalization in the Insect Gut

Begin with a chemically fixed gut excised from an insect carrying a plant virus.
Wash with a buffer to remove the fixative.
Add a non-ionic detergent to permeabilize the cellular membranes.
Wash again to remove any detergent residues.
Add a solution containing two different fluorescent antibodies and a blocking agent.
Upon incubation, the blocking agent saturates nonspecific binding sites, while the antibodies enter the cytoplasm.
One antibody binds to the viral antigen within the host cell vesicles, and the other binds to a vesicle-associated membrane protein.
Wash to remove the unbound antibodies.
Add a fluorescent dye that binds to the actin filaments to visualize the host cells.
Wash to eliminate any unbound dye.
Mount the stained gut in a medium containing a fluorescent nuclear stain to label the cellular DNA.
Using confocal microscopy, visualize the coexistence of the green and red fluorescent signals, indicating colocalization of the viral antigen and the membrane protein.

Prepare the antibodies and mounting medium as described in the text manuscript.

Place the freshly excised and PBS washed WBPH guts in 100 microliters of 4% paraformaldehyde in a 200 microliter centrifuge tube at room temperature. After two hours, replace the paraformaldehyde solution with 200 microliters of PBS. After 10 minutes, remove the PBS to eliminate any paraformaldehyde and repeat the PBS wash step twice.

After two PBS washes, add 200 microliters of non-ionic detergent Triton X-100 to permeabilize the samples at room temperature. After 30 minutes, remove the Triton X-100 and wash away any remaining detergent with three 10-minute washes with PBS. Dilute the labeled antibodies one to 50 with 50 microliters of bull serum albumin.

Add the diluted antibodies to the tube and incubate the samples overnight at four degrees Celsius. On the morning after removing the antibody, wash away the remaining antibody diluent with three 10-minute washes with PBS. Dilute one microliter of Dylight 633-Phalloidin with 50 microliters of PBS and add 50 microliters of diluted phalloidin to the tube for a two-hour incubation at room temperature.

After removing phalloidin, thoroughly wash away the remaining phalloidin with three 10-minute washes with PBS. Place a drop of mounting medium containing DAPI on a microscope slide. Transfer the guts to the medium and gently place the cover glass over the sample without creating bubbles.