This protocol can be used to study various movement in insects, functions of insect proteins, and interactions between virus and vector insect in vivo. This method is efficient and informative. The structures of the insect gut and the auxiliary cells are clearly visualized when viewed with the laser scanning confocal microscopy.
Demonstrating the procedure will be Lu Zhang, PhD from our laboratory. To begin, transfer non-viruliferous insects from glass beakers onto fresh southern rice black streaked dwarf virus, SRBSDV, infected rice plants covered with an insect proof net for a two-day virus acquisition access period. Then collect the insects in glass beakers containing fresh rice seedlings.
After two days, collect the insects from the glass beakers using a manual aspirator for dissection and excision of the gut. Prepare 0.01 molar PBS, 4%paraformaldehyde, and 2%Triton X-100 as described in the text manuscript. Use a pipetter to place 100 microliters of PBS on a glass slide and place the slide on the stage of an optical microscope.
Use manual aspirator to collect the SRBSDV infected adults from the glass beakers and place them in a 1.5 milliliter tube on ice to paralyze the insects. Transfer a paralyzed adult into the 100 microliters of PBS on the slide with the abdomen up. Use tweezers to clamp the body and remove the head with another set of tweezers.
With one set of tweezers, clamp the sides of the abdomen and clamp the ovipositor or the copulatory organ of the tail with the other set, then carefully pull the intersegmental membrane of one abdominal segment to slowly expose the gut in the abdomen. Continue tearing away the membrane and gradually pull out the complete gut from the abdomen. Gently pull off the tail which is connected to the end of the gut to remove the complete gut.
Place excised guts into a 200 microliter centrifuge tube. Add 200 microliters of PBS to the tube and use a pipette to thoroughly wash the guts. Use a pipetter to place 100 microliters of PBS on a glass slide and place the slide on the stage of an optical microscope.
Use manual aspirator to collect the SRBSDV infected nymphs from glass beakers and place them in 1.5 milliliter tube placed on ice to paralyze the insects. Transfer a paralyzed nymph into the 100 microliters of buffer on the slide with the abdomen facing up. Detach the tail of the nymph, then clamp the insect body to fix it gently and use the other pair to clamp the head.
Gently pull the head away from the body while still maintaining its attachment to the gut so that the head is detached from the body, but the gut is still attached to the thorax and abdomen. With the tweezers still clamping the body, use the other pair to move the head carefully and gradually pull out the gut. Detach the gut from the head to obtain an intact gut without damaging the body.
Place the excised guts into a tube. Add PBS to the tube and gently suck release the solution with a pipette to wash the guts thoroughly. 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.
The representative laser scanning confocal micrograph of excised WBPH guts from adults after labeling with phalloidin showed three sections:the foregut, midgut, and hindgut. Among these, midgut is the initial infection site of SRBSDV. The monolayer epithelial cell structure of the gut facilitates the study of the cellular localization of insect proteins and colocalization of viral and insect proteins.
Excised WBPH guts with DyLight 488 labeled anti-VAMP-7 and SRBSDV virions with DyLight 549 labeled anti-SRBSDV antibody are shown here. VAMP-7 and SRBSDV virions were shown to co-localize in the cytoplasm suggesting that VAMP-7 may play a role in virus transmission in vivo. Permeabilized guts should be cleaned thoroughly after incubation with luciferin conjugated antibodies to reduce the background and non-specific binding.
This is a reliable method to view the localization of virus, other pathogens, and insect proteins in insects. It provides the foundation for further studies of the relationship between pathogens and insects.
