Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Whole-mount staining of Drosophila tissues, such as testis samples, is often problematic due to limited antibody penetration. Our fixation protocol using NP40 and heptane permits a uniform and reproducible labeling. By maintaining tissue shape while permitting deep antibody penetration, this procedure facilitates the reproducible, quantifiable acquisition of immunostaining in three dimensions, both singly and in colocalization studies.

For diploid germ cell evaluation, zero to 15 hours post-eclosion, anesthetize the flies by a brief exposure to carbon dioxide, and transfer the flies to a fly pad. Place the pad under a dissecting microscope at a 10X magnification, and use forceps to remove the heads from five to 10 male flies. Transfer the decapitated fly bodies to 100 microliters of Ringer's buffer on a silicone-coated glass slide on a black background support, and use the 40X magnification and a pair of number five forceps to grasp a fly between the thorax and the abdomen.

Then pull the abdomen away from the thorax and rapidly isolate the testes and adjacent tissues from the fly carcass. Once all of the testes have been collected, clean the samples of any remaining tissue and use a 200 microliter pipette tip to remove the excess buffer from around the dissected testes. Then place a drop of 4%paraformaldehyde onto the testes and quickly transfer the tissues to a tube containing two milliliters of freshly prepared 4%paraformaldehyde solution supplemented with 20%NP40 and heptane.

For thorough fixation of the testes samples, shake the tube vertically for 30 seconds before incubating the testes on a rotary mixer at room temperature for 30 minutes. At the end of the incubation, transfer the tube to a tube rack and allow the testes to settle at the bottom of the tube. Remove all of the heptane and fixative solution and quickly rinse the testes three times with PBS.

After the last wash, transfer the testes to a new 200 microliter tube and use a 200 microliter pipette equipped with a 10 microliter pipette tip to remove any excess buffer. Then add 200 microliters of fresh PBS-To the tube and place the testes on ice. For antibody labeling of the testes samples, replace the PBS with 200 microliters of 0.3%PBS-T for a 30-minute incubation at room temperature.

After permeabilization, block any non-specific binding with 0.3%PBS-T plus 5%normal goat serum for one hour at room temperature before adding 200 microliters of the primary antibody of interest diluted in 0.3%PBS-T plus 5%normal goat serum for an overnight incubation at four degrees Celsius or room temperature with rocking. The next morning, wash the samples with three 15-minute washes with fresh 0.3%PBS-T and rocking at room temperature per wash and allow the testes to settle to the bottom of the tube by gravity. Add 200 microliters of an appropriate secondary antibody to the tube for a four-hour incubation with rocking at room temperature protected from light before washing the samples in fresh 0.3%PBS-T per wash, as demonstrated.

After the 0.3%PBS-T wash, wash the samples one time in 0.2%PBS-T and one time in 0.1%PBS-T for 30 minutes per wash on the rocker, followed by a final 30-minute wash in PBS alone. After the PBS wash, label the samples with 180 microliters of a one microgram per milliliter DAPI solution in PBS for 15 minutes at room temperature protected from light, followed by three 5-minute washes in fresh PBS per wash. After the last wash, use a hydrophobic barrier pen to draw a circle on a clean slide, and use a 0.1%PBS-T primed pipette tip to transfer the testes to the slide.

Aspirate any excess PBS and add 100 microliters of Citifluor AF1. Gently place a glass cover slip over the tissue, taking care to avoid trapping air bubbles, and use clear nail polish to seal the cover slip to the slide before observing the slides by confocal fluorescence microscopy. After acquiring confocal images of the samples, import the images into an appropriate image analysis software program.

Then use manual segmentation to define the nucleus within each cell and determine the Pearson's correlation coefficients inside the entire volume between two fluorophores. This protocol permits the acquisition of confocal images of a sufficient quality to be treated for 3D reconstitution. For example, in these four adjacent serial sections of a stage S5 spermatocyte, a low level of Mad1 is observed at the nuclear envelope and a high-level accumulation of Mad1 can be visualized in conjunction with DAPI staining within the nucleus.

in this 3D reconstituted image, the low level presence of Mad1 at the nuclear surface and high-level accumulation of Mad1 within the nucleus can be observed together with the DNA. In these 3D reconstitutions, large S5 spermatocytes stained with Nucleoporin 62 and Mad1 or Importin-beta and DAPI can be observed. In both reconstructions, the nuclear pore-associated components appear to be non-uniformly distributed over the nuclear rim as dots of irregular size.

While conserving the 3D volume of the cells, this protocol also facilitates a uniform penetration of antibodies of interest into the depth of every cell, allowing quantitative analyses, such as colocalization. Because the protocol permits a reliable measurement of colocalizations from independent testes, it can also be used to compare staining in different fly genotypes. The amount of NP40 used is critical for sufficient permeabilization.

If the procedure is followed closely, reproducible labeling and 3D nuclear structure preservation can be achieved. Although whole-mount staining of tissues within Drosophila can be tricky, this protocol can be easily adapted for other samples.