Dissection of Drosophila melanogaster Indirect Flight Muscles for Microscopy Approaches

We study muscle development, in particular, how RNA regulation contributes to sarcomere formation and fiber type-specific function in Drosophila melanogaster. The indirect flight muscles are assembled during pupal development. To follow this process in vivo requires developing and optimizing technically demanding dissection protocols.

To begin, position a microscope slide under a stereo dissecting microscope. Use a plastic pipette to transfer a drop of PBS to the slide. With a paintbrush or a pair of forceps, transfer sample Drosophila flies in small groups to the drop of PBS.

Then, use a pair of Vannas spring scissors to remove the head, wings, and abdomen, while leaving the legs attached. Now, gently transfer thoraces to 500 microliters of fixation solution in one well of a 24-well plate with the help of a brush. Place the plate on a nutator or rocking shaker and fix for the desired duration, typically between 15 to 60 minutes.

With a P1000 microliter pipette, remove the buffer from the well. Place a drop of 0.05%PBS with Triton X-100 on a microscope slide under a stereo dissecting microscope. Then, use a brush or forceps to transfer the thoraces to the PBST drop.

Orient the thorax with the scutellum pointing upwards and stabilize it between a pair of Dumont 3 forceps. Next, slide a Cryostat blade across the thorax to notch the scutellum. Then, cut down in one smooth, downward stroke along the midline to create the thorax hemisection and expose the indirect flight muscles.

Using forceps, transfer thorax hemisections from the microscope slide to 500 microliters of 0.05%PBST into one well of a 24-well plate for further processing. To perform dissection of pupal indirect flight muscles, collect the pre-pupae age to the correct time point on a wetted filter paper. For pupal hemithorax dissections, place a strip of double-stick tape adhered to a microscope slide under a stereo dissecting microscope at 10X to 20X magnification.

At the desired time point, use a paintbrush to transfer pupae from the filter paper to the tape. Use a pair of 5 forceps to open the front of the pupal case and gently remove the operculum. Carefully slit along the dorsal side of the pupal case and peel it away in 1 to 2 millimeter increments.

Once freed from the pupal case, gently transfer the pupa to a drop of PBS on a second microscope slide. Now, use Vannas spring scissors to remove the abdomen. Using a brush or forceps, transfer the head and thorax to 500 microliters of fixation solution into a well of a 24-well plate.

After fixation and removal of the fixation solution, transfer the thoraces to a drop of 0.05%PBST on a microscope slide. Separate one thorax from the rest and orient it ventral side down, stabilized between Dumont 3 forceps. Slide a Cryostat blade across the thorax until the basal membrane and cuticle are slit open.

Then, cut down in one smooth, downward stroke along the midline to generate thorax hemisections and expose the indirect flight muscles. Use a paintbrush or forceps to transfer hemisections to 500 microliters of 0.05%PBST in a 24-well plate for further staining. For open book dissection of the indirect flight muscles, transfer pupae to a black silicone dissecting dish filled with PBS after removing them from the pupal case.

Using Dumont 5 forceps, gently press the dissected pupae to the dish surface and insert two insect needles through the abdomen of each pupa to secure them in a line. Using Vannas spring scissors, cut open the basal membrane and anterior head region of each pupa. Now, insert the scissors into the opening and cut along the sides of the pupa.

Use forceps to lift and remove the ventral portion with scissors. Then, remove the brain, ventral nerve cord, trachea, and gut. Use a 200 microliter pipette with a clipped tip to gently pipette PBS over the thorax and remove remaining fat bodies.

Then, use a pair of Vannas spring scissors to cut down the midline of the dorsal thorax to create two leaflets containing indirect flight muscles. Cut the leaflets of the dorsal thorax away from the abdomen and transfer them to 300 microliters of fixative solution in a black glass dish using Dumont 5 forceps. After fixing for the appropriate time, use a pipette to remove fixative and wash in 500 microliters of 0.05%PBST before immunostaining.

Once immunostaining is complete, pipette out the secondary or stain mixture. Then, wash the samples four times for 10 minutes each in 0.05%phosphate-buffered saline with Triton X-100 at room temperature. Label the frosted area of a microscope slide with the sample number or identifying information, such as date, genotype, antibody, or slide number.

Pipette a drop of glycerol on the microscope slide to secure spacers. Then, position 1 coverslip spacers on the microscope slide about 1 centimeter apart. Now, add a drop of mounting medium between the spacers.

Pipette out any residual wash buffer, and immediately transfer samples into the mounting medium. Using Dumont 5 forceps, organize the samples into non-overlapping rows and columns with the indirect flight muscles facing up. Carefully add a 1 coverslip over the samples.

Next, backfill the sample area with mounting medium until all thoraces are enclosed. Then, use clear nail polish to seal both the sample coverslip and spacer coverslips. Sarcomere morphology was effectively preserved with 4%paraformaldehyde, but a significant difference in sarcomere width was seen between paraformaldehyde and PBS compared to relaxing solution.

Methanol fixation preserved myofiber and sarcomere structures, but sarcomeres were significantly shorter and thinner than with 4%PFA fixation. Glyoxal fixation significantly reduced sarcomere width compared to 4%paraformaldehyde. Fixation for 15 or 30 minutes preserved sarcomere structure, but a seven-minute fixation produced inconsistent morphologies.

The myofibril structure of the indirect flight muscles appears frayed and ripped due to dull blades or blade slips. Forceps in direct contact with the muscles led to divots or stretching, pulling the sarcomeres apart. Stretching or thorax deformation led to a loss of the Z-disc.

Sawing the fibers caused unraveling. At 72 hours after puparium formation and adult stages, SmnE33 mutants lacked sarcomeres, with reduced F-actin and starburst bundles. At 26 hours after puparium formation, SmnE33 and controls showed similar SmnE33 cables and mesh networks.

We received protocols for immunofluorescence microscopy of indirect flight muscles as early as 16 hours after puparium formation, all the way through late pupal development and adult stages. Our protocol enables us to look at molecular phenotypes and cellular deficits as they pertain to protein localization and muscle structure and function that usually result in behavioral deficits.