Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

The overall goal of this procedure is to visualize or sate tissues in C two using fluorescence microscopy following sonication. This is accomplished by first collecting embryonic and larval samples and then fixing them using formaldehyde, heptane, and methanol. The second step is to sonicate the fixed sample in order to remove the cuticle to permit antibody penetration.

Next, the sample is immuno stain with primary and fluorescent secondary antibodies. The final step is to mount the sample in a glycerol based Antifa solution. Ultimately, confocal or epi fluorescent microscopy is used to visualize the developing tissue as well as protein expression and localization.

The main advantage of this technique over other methods like tissue dissection, is that it allows for in visualization of a variety of developing tissues. Generally, individuals new to this method will struggle because the sonication step is dependent on a number of factors to achieve maximum efficiency, including the proper positioning of the sonica probe. Demonstrating the procedure will be Ashley Fiddler and Lauren Bole, two talented undergraduate researchers from my lab.

To begin the experiment, carefully remove embryos collected on an agar plate using a small paint brush wedded with phosphate buffer Triton X 100 or PBTX solution. Next, wipe the sample laden paintbrush against the interior facing ridge of a cell strainer. Place a Petri dish cover underneath the cell strainer.

Then rinse the paintbrush and the walls of the cell strainer with a squirt bottle containing PBTX. After the sample has been transferred to the strainer, pour enough PBTX into the strainer to raise the sample off the mesh. Repeat this step three times to remove yeast and fly waste, remove PBTX.

Then pour 50%bleach solution into the strainer and allow the larvae to sit in the solution for five minutes. Next, remove the solution. Then wash the sample with PBTX and allow the sample to sit in the solution for three minutes.

Repeat the step five times, dry the bottom of the cell strainer. Then use a paintbrush wedded with PBTX to transfer the sample into a scintillation vial containing 1.75 milliliters of pems. In a fume hood, add 250 microliters of 37%formaldehyde and eight milliliters of reagent grade.

Heptane to the scintillation vial, and shake the vial at 200 RPM for 20 minutes. Then add 10 milliliters of methanol to the scintillation vial without removing the aqueous phase and shake vigorously at 500 RPM. After shaking immediately pour the contents into a cell strainer over a liquid waste container.

Add extra methanol to the vial and run the contents through the cell strainer. To ensure the entire sample has been removed. Try the bottom of the cell strainer with laboratory tissue.

Then apply a paintbrush to transfer the sample from the strainer to a 1.5 milliliter micro centrifuge tube containing 0.5 milliliters of methanol. Wait for the sample to settle. Then remove the methanol with a pipette and replace it with a fresh 0.5 milliliter aliquot of methanol.

Repeat this step three times and add a final 0.5 milliliter of methanol to the tube before storing it at negative 20 degrees Celsius. For later, use retrieve the stored sample from the freezer and remove the methanol with a pipette. Then add one milliliter of a 50%methanol phosphate buffer tween or PBTW solution to the tube, and rock the tube for three minutes.

Next, rinse the sample with one milliliter of PBTW twice allowing the sample to settle before drawing off the PBTW with a pipette, add 1.0 milliliters of bovine serum albumin phosphate buffered tween, or BBTW to the vial, and rock it for three minutes. Allow the sample to settle and remove the BBTW with a pipette. Repeat this Step two times.

Taking care to remove as much BBTW as possible without losing any of the sample. Then add 0.5 milliliters of BBTW to the tube and place it on ice. Next, set the sonicate to 10%maximum amplitude and a runtime of two seconds with constant ation, clean the ator probe by placing the probe tip into a 50 milliliter conical tube filled with deionized water and run the sonicate.

Then dry the sonicate probe with a laboratory tissue. Submerge the probe into the tube, three to four millimeters above the sample and start the sonication. Then remove the tube and place it on ice for 30 seconds to prevent overheating from the sonication, and to allow the sample to settle at the bottom of the tube.

Repeat this step as required by sample age after sonication is complete. Rinse the sample two times with one milliliter of BBTW. After each rinse, allow the sample to settle before drying off BBTW With a pipette, add one milliliter of BBTW to the vial and place it on a rocker.

After three minutes, allow the sample to settle and remove the BBTW with a pipette. Then repeat this. Step two times, block the sample with 0.5 milliliters of 5%normal serum diluted in BBTW to the tube, and place the sample on a rocker for one hour at room temperature.

Next, add 0.5 milliliters of primary antibody solution and rock the sample overnight at four degrees Celsius, allow the sample to settle to the bottom of the tube. Then draw off primary antibodies with a pipette and rinse twice with one milliliter of BBTW. Add one milliliter of BBTW to the vial and then place it on a rocker for three minutes.

Then remove the BBTW with a pipette and repeat this. Step five times, block the sample with 5%normal serum BBTW solution and rock it for 30 minutes. Then remove the solution.

Add secondary antibody diluted at 5%normal serum BBTW solution and wrap the tube in aluminum foil. Then rock the sample overnight at four degrees Celsius. Draw off secondary antibodies with a pipette and rinse the sample with one milliliter of PBTW twice.

Next, add one milliliter of PBTW to the vial and place it on a rocker for five minutes. Then allow the sample to settle and remove the PBTW with a pipette. Repeat this.

Step three times. Add 80 to 100 microliters of DAB co solution to the tube and store it at negative 20 degrees Celsius. Finally, to mount the sample, use five to 10 microliters of dab BCO p lene diamine, or PPD anti fade.

Then view it with fluorescence microscopy. These Z projections of four confocal slices demonstrate the efficacy of the protocol for visualizing morphological features as well as individual cells in C two during late embryonic through early and mid L three stages of drosophila development. Testes development is a particularly good system for illustrating protocol efficacy because testes maturation is dynamic throughout larval development.

These images highlight germ cells in red and hub cells and FU zones in green single confocal sections of mid L three brains. Immuno stain for ganglion mother cells, undifferentiated neurons in green and immature and primary neurons. In red show strong staining with distinct expression patterns in the brain lobes and the ventral nerve cord, depending on mounting orientation, imaging allows for brain tissue visualization from the dorsal surface, ventral surface, or in sagittal cross-section.

This in situ two technique allows biologists to explore organ development in tissue morphogenesis in organisms where protective cuticle prevents antibody penetration. After watching this video, you should have a good understanding of how to sonicate and immunostain late stage embryonic and larval esophagus samples in order to visualize developing tissues in C two. Don’t forget that working with formaldehyde, heptane and methanol can be extremely hazardous and precautions such as working in a fume hood.

Wearing appropriate gloves and a lab coat should always be observed while performing this procedure.