Dissection of Drosophila melanogaster Indirect Flight Muscles for Microscopy Approaches

Animals have hundreds of muscles that control different movements, from enabling ambulation to facilitating speech to grasping a tool. To support such a wide array of function, muscles differ in size, attachment, contractile ability, metabolism, and gene expression¹. A major focus within the field of muscle biology is understanding how these structural and functional differences arise during development and are altered in muscle disease². Muscle contraction is powered by sarcomeres built with actin-containing thin filaments anchored at the Z-disc that interdigitate with bipolar myosin-containing thick filaments anchored at the M-line³. Sarcomeres are organized end-to-end into myofibrils, and this regular arrangement in striated muscle appears as alternating light and dark bands under a microscope⁴, providing a practical visual method to monitor changes in sarcomere assembly and organization. In combination with genetics and biochemical approaches, microscopy has provided important insights into cellular processes such as transcriptional regulation, RNA processing, metabolism, and mechanotransduction^5,6,7 that instruct and fine-tune muscle development and function. Many of these cellular processes are disrupted in human myopathies and neuromuscular disorders^8,9, underscoring the importance of understanding muscle development to gain insight into the etiology of muscle disease. Advances in genetic engineering, enabling the manipulation of individual cells and endogenous labeling of proteins^10,11 coupled to technological advances in image acquisition and resolution^12,13 ensure the continuing importance of microscopy to muscle research. Model systems with clear and easily implemented protocols for muscle dissection and sample preparation are thus valuable in the muscle field.

The indirect flight muscles (IFMs) of D. melanogaster are a powerful and well-characterized model to study basic principles of myogenesis⁸. IFMs consist of six dorsal-longitudinal muscles (DLMs) per hemithorax that form on larval muscle templates and seven dorsal-ventral muscle fibers (DVMs) that are built de novo during pupal development¹⁴. Due to their accessibility near the midline, many IFM studies focus on the DLMs, the largest muscle in adult flies spanning the entire 1 mm length of the thorax¹⁵. At the onset of pupation, IFM myoblasts migrate from their niche on the wing disc hinge to the thorax, where they fuse to form syncytial myotubes¹⁶. After tendon attachment around 24 h APF at 27 °C, IFM myofibers progressively organize their cytoplasm and cytoskeleton, compact, and initiate myofibrillogenesis from 30-32 h APF¹⁷. New sarcomeres are added to myofibrils and myofibers grow to fill the thorax until around 48 h APF. IFMs then undergo a maturation process concordant with changes in both transcription and splicing that promotes the acquisition of adult metabolic and contractile properties and stretch activation¹⁸. Adult flies eclose from the pupal case between 90 h and 100 h APF at 27 °C, and at approximately 100 h APF (females at about 98 h APF and males at about 102 h APF) at 25 °C¹⁹. IFMs are a versatile and relevant model system. They can be monitored at all stages of myogenesis, are morphologically and functionally distinct from other myofiber types in the fly, and benefit from a wide assortment of accessible genetic tools and manipulation approaches^8,20. The contractile mechanism, basic sarcomere morphology, and many structural components are conserved from flies to vertebrates⁵, enabling translation of myogenic principles from flies to humans. Indeed, fly models of neuromuscular diseases such as myotonic dystrophy, spinal motor atrophy (SMA), myofibrillar myopathies, and actinopathies have provided etiological and therapeutic insights^21,22. Notably, flies are well-suited for microscopy to analyze myofiber and sarcomere morphology across myogenesis, from myoblast specification to myofibrillogenesis to sarcomere maturation and maintenance.

The Drosophila IFMs have been used for microscopy applications for more than four decades, greatly contributing to the understanding of muscle growth and development⁸. During this time, IFM dissection protocols have been continuously adapted as technology advanced, leading to a wide array of protocols presented in variable levels of detail that are optimal for some and suboptimal for other imaging techniques. For example, protocols developed for electron microscopy involve strong fixation conditions, ultrathin sectioning, and contrast staining with heavy metals^3,23,24,25, but are not optimal for light microscopy. Protocols developed to rapidly dissect IFMs out of the thorax for use in biochemical assays or molecular analyses^26,27 lose the tissue context and can disrupt myofibril structure, and are therefore suboptimal for microscopy approaches. Dissection approaches were developed for analysis of larval muscle or adult abdominal muscle^20,28, but are not optimal for isolation of intact IFMs. Live imaging approaches require either minimally invasive dissections or special media to keep IFM tissue alive during the imaging process^29,30, and do not involve the fixation steps necessary to preserve muscles for immunohistochemistry. Multiple protocols exist for paraffin-embedding or cryo-sectioning of IFM tissues and are compatible with histochemistry or antibody staining^27,31,32, but these approaches typically involve freezing and tissue dehydration that can alter muscle ultrastructure and limit recognition of certain antigens by primary antibodies. Protocols that directly fix the IFMs preserve muscle morphology and limit changes in antigen conformation and are desirable for many immunohistochemistry experiments²⁴. However, these protocols are also targeted to specific applications, for example, protocols that dissociate IFMs enabling single-myofibril analysis by fluorescence, cryoEM, or superresolution microscopy^20,33,34. Other examples include protocols to dissect intact IFM fibers from the thorax^32,35 or to generate whole-mount or hemithorax preparations^20,32,36 that enable analysis of both myofibers and sarcomeres in an intact context. It can be challenging for a new user to sort through the large number of existing protocols, the majority of which are text-based. Trainees and undergraduate researchers would benefit from a detailed IFM dissection protocol as applied to standard immunohistochemistry applications.

This article includes a series of protocols for the dissection and preparation of D. melanogaster IFMs at pupal and adult stages for immunohistochemistry microscopy approaches. This approach has been used in recent publications tracing developmental functions of the RNA-binding proteins Bruno1³⁷ and Rbfox1³⁸ throughout IFM myogenesis. The protocol includes detailed illustrations and step-by-step images to guide potential users through IFM hemithorax dissection of adult and late pupal (48 h to 96 h APF) IFMs, where the thorax is bisected along the midline, keeping myofiber structure intact. In addition, this protocol covers open-book dissection of early pupal (8 h to 48 h APF) stages, where the ventral portion of the thorax is slit and lifted, like opening a book, to expose the intact IFMs attached to the dorsal epidermis. Further steps in the protocol include fixation, staining, and sample mounting, and representative data demonstrate that this approach is compatible with diverse fixation methods to prepare tissues for immunohistochemistry and fluorescence microscopy. Representative examples are provided to aid new users in identifying common dissection artifacts. The protocol is applied to analyze the developmental IFM phenotype of Smn^E33, a hypomorphic allele of survival motor neuron (Smn) used as a fly model of Spinal Motor Atrophy (SMA)^21,39, at 26 h APF, 72 h APF, and in adult IFMs. The data presented here provides insight into the muscle-specific function of Smn and highlights the applicability of this dissection protocol to study muscle development and the etiology of muscle disease.

Ethical statement:
All experiments in this protocol use Drosophila melanogaster, a holometabolous insect. Invertebrates are not subject to animal welfare regulations in the United States of America, and their use does not require ethical approval. All work was conducted under standard institutional biosafety and laboratory safety guidelines, and was approved by the University of Missouri Kansas City Institutional Biosafety Committee under protocol number 21940 (22-11).

1.Hemithorax dissection of adult IFM (Figure 1)

1. Assemble the necessary supplies, including two Dumont #5 forceps with biology tips, a Dumont #3 forceps, a pair of Vannas spring scissors, a plastic pipette, a paint brush, a 24-well plate, lint-free wipes, a microscope slide, and cryostat blades (see Table of Materials). Prepare 1x PBS, 0.05% PBS-T, and fixation solutions (see Supplementary File 1). A flow-diagram overview of the following adult hemithorax dissection is available (Supplementary Figure 1).
2. Collect adults of the desired genotype and age (for example, newly eclosed, 1 day adult, or 5 day adult). Anesthetize sample flies on CO₂ or ice.
   NOTE: Flies used in this protocol were grown on a 12-hour day: 12-hour night cycle in a temperature and humidity-controlled incubator. Drosophila are standardly grown at 25 °C or 27 °C, and can be grown at 18 °C, which extends the growth cycle from 10 days to approximately 20 days.
3. Use a plastic pipette to transfer a drop of 1x PBS⁴⁰ to a microscope slide under a stereo dissecting microscope.
4. Transfer the sample flies to the PBS drop in small groups using a paint brush or forceps (Figure 1A). Remove the head (Figure 1B), wings (Figure 1C - D), and abdomen (Figure 1E) using a Vannas spring scissors. Leave the legs attached (Figure 1F).
5. Gently transfer thoraces using a brush or forceps to 500 µL fixation solution in one well of a 24-well plate. Fix for the desired length of time (typically 15-60 min) on a nutator/rocking shaker.
   NOTE: This protocol is compatible with multiple fixation buffers, including paraformaldehyde, glutaraldehyde, methanol, glyoxal solution, and others. See Representative Results and Supplementary File 1 for details on optimizing fixation conditions. Fixatives should be handled using gloves and appropriate personal protective equipment (PPE) according to your institution's hazardous chemical guidelines.
6. Remove fixative with a 200 µL pipette and wash in 1 mL of 0.05% PBS-T on a nutator/rocking shaker for 5 min at room temperature.
   NOTE: Dispose of fixative following the institution's hazardous chemical guidelines. For most antigens, fixed thoraces can be stored in 1 mL of 0.05% PBS-T for up to 2 weeks at 4 °C prior to cutting and staining. Higher concentrations of Triton X-100 should be tested before use, as long incubations in detergent can degrade sarcomere structure.
7. Remove buffer with a 200 µL pipette and transfer thoraces using a brush or forceps to a drop of 0.05% PBS-T on a microscope slide under a stereo dissecting microscope.
8. Orient the thorax with the scutellum pointing up and stabilized between a pair of Dumont #3 forceps (Figure 1G-H).
9. Slide a cryostat blade across the thorax to notch the scutellum (Figure 1H, J, L), and then cut down in one smooth downward stroke along the midline (Figure 1I, J, L - N) to generate the thorax hemi-section exposing the IFMs (Figure 1K).
   NOTE: It is important that the cuticle is "notched" or cracked prior to cutting through the IFMs to avoid stretching artifacts that damage sarcomere structure. Do not "saw" the blade back and forth, as this motion mechanically damages the IFMs, resulting in frayed myofibrils. Too much buffer destabilizes the thorax between the forceps and makes the cut more difficult. Alternate approaches include stabilizing the thoraces one at a time on double-stick tape to facilitate cuts, as well as cutting from the ventral side of the thorax.
10. Move the two halves of the thorax to a position on the slide where they will remain covered in buffer to avoid drying out, but are not in the way of future cuts. Repeat steps 1.8-1.10 until all thoraces have been cut.
11. Use a forceps to transfer thorax hemi-sections from the microscope slide to 500 µL of 0.05% PBS-T in one well of a 24-well plate and proceed with blocking and immunostaining (see section 4 below).

2. Hemithorax dissection of late pupal IFM (~48 h to 96 h APF) (Figure 2, Figure 3)

1. Stage pupae of the desired genotype and age to the selected point in pupal development (Figure 2). Collect male or female white pre-pupae (Figure 2A, B) using a damp paintbrush and incubate on wetted filter paper in a 60 mm cell culture dish. Pupal staging and removal from the pupal case has been described previously^20,26.
   NOTE: Pupae undergo multiple morphological changes as they age, such as head eversion and pigmentation of the pupal case, eyes, and bristles^19,41 (Figure 2C). These morphological features can be used as a rough guide to identify the age of a pupa (Figure 2D-E) and to identify pupae that fail to develop normally (Figure 2F). However, to ensure an accurate staging timeline, it is recommended to collect white pre-pupae in narrow time windows, for example staging every 30 min or every 2 h. Tightly-staged timepoints are necessary to distinguish major changes in cytoskeletal structure and myofiber length that occur from 24 h to 35 h APF, as well as to distinguish differences in sarcomere structure at 35h, 48 h, 60 h, 72 h, 80 h, and 90 h APF^18,42.
2. Assemble the necessary supplies, including a paint brush, two Dumont #5 forceps with biology tips, a pair of Vannas spring scissors, double-stick tape, a microscope slide, a plastic pipette, a 24-well plate, Kimwipes, and cryostat blades (see Table of Materials). Prepare 1x PBS, 0.05% PBS-T, and fixation solutions (see Supplementary File 1). A flow-diagram overview of the following pupal hemithorax dissection is available (Supplementary Figure 2).
3. At the desired time point, use a paintbrush to transfer pupae from the filter paper to a strip of double-stick tape adhered to a microscope slide under a stereo dissecting microscope at 10-20x magnification (Figure 3A,B). Line up the pupae on the tape facing the same direction, ventral side down.
4. Use a pair of #5 forceps to open the front of the pupal case of the first pupa, gently removing the operculum (the region over the head) (Figure 3C, D, I).
5. Carefully use a #5 forceps to slit along the pupal case just dorsal to the wing and peel the pupal case away from the pupa in 1-2 mm increments. Affix the pupal case to the double-stick tape as one works from anterior to posterior (Figure 3E - G, I). Avoid poking the pupa itself, rather insert the forceps tip between the pupal case and the basal membrane/pupal sac surrounding the pupa.
6. After the case has been removed, use a scooping motion with the forceps to lift under (Figure 3H, I) and gently transfer the pupa out of the case to a drop of 1x PBS on a second microscope slide (Figure 3J, K) to avoid dehydration. Repeat steps 2.4-2.6 (Figure 3I) until all pupae have been removed from the pupal case.
7. Remove the abdomen from the dissected pupae using Vannas spring scissors (Figure 3L).
   NOTE: The abdomen is removed to enable better penetration by fixative in the following step. To avoid damaging the thorax, cut directly in the abdomen, leaving up to 30% of the abdomen attached to the thorax.
8. Gently transfer the pupal head with the thorax (Figure 3M) using a brush or forceps to 500 µL of fixation solution in one well of a 24-well plate. Fix for the desired length of time (typically 15-60 min) and wash in 1 mL of 0.05% PBS-T as described in steps 1.5-1.6.
9. Remove buffer with a 200 µL pipette and transfer thoraces using a brush or forceps to a drop of 0.05% PBS-T on a microscope slide under a stereo dissecting microscope under 10x magnification.
10. Separate one sample from the rest and orient it with the ventral side down and stabilized between a pair of Dumont #3 forceps (Figure 3N).
11. Slide a cryostat blade across the thorax just until the basal membrane and cuticle are slit open, and then cut down in one smooth downward stroke along the midline (Figure 3O) to generate the thorax hemi-sections exposing the IFMs (Figure 3P).
    NOTE: Do not "saw" the blade back and forth, as this motion mechanically damages the IFMs resulting in frayed myofibrils. Too much buffer destabilizes the thorax between the forceps and makes the cut more difficult. Pupal thoraces can be cut with the head attached and oriented either towards the left or the right.
12. Move the two halves of the thorax to a position on the slide where they will remain covered in buffer to avoid drying out, but are not in the way of future cuts. Repeat steps 2.10-2.12 until all thoraces have been cut.
13. Use a forceps to transfer thorax hemi-sections from the microscope slide to 500 µL of 0.05% PBS-T in one well of a 24-well plate and proceed with blocking and immunostaining (see step 4 below).

3. Open-book dissection of early pupal IFM (~15 h to 48 h APF) (Figure 4)

1. Stage pupae of the desired genotype and age to the selected point in pupal development (Figure 2) as described above (Step 2.1).
   NOTE: For open-book dissections, pupae should be dissected in groups of no more than 8-10 flies.
2. Assemble the necessary supplies, including a paint brush, two Dumont #5 forceps with biology tips, a pair of Vannas spring scissors, double-stick tape, a microscope slide, a plastic pipette, a black silicon dissecting dish, a black glass staining dish, insect pins, and lint-free wipes (see Table of Materials). Prepare 1x PBS, 0.05% PBS-T, and fixation solutions (see Supplementary File 1). A flow-diagram overview of the following pupal open-book dissection is available (Supplementary Figure 3).
3. At the desired time point, remove pupae from the pupal case as described in steps 2.3-2.6. Transfer pupae to a black silicon dissecting dish filled with 1x PBS after removal from the pupal case (Figure 4A).
4. After all pupae are freed from the pupal case, use a Dumont #5 forceps to overcome surface tension and gently push dissected pupae down to the surface of the silicon. Arrange the pupae in a line near the center of the dish (Figure 4B).
5. Insert two insect needles using a Dumont #5 forceps through the abdomen of each pupa to hold it in place (Figure 4C,D). Ensure that the pupae are oriented with their ventral side up.
6. Use a Vannas spring scissors to cut open the basal membrane and anterior head-region of each pupa (Figure 4E).
7. Insert the scissors into the opening and cut along the right and left sides of each pupa (Figure 4F, G, Q). Position the scissors and cut just below the developing notum back to the junction between the thorax and abdomen to remove the legs and wings with the ventral portion. Ensure the scissors are inserted horizontally and do not contact the dorsal portion of the thorax (Figure 4Q).
8. Use a Dumont #5 forceps to grasp and lift the ventral portion of the pupa and remove it with a Vannas spring scissors (Figure 4H, I, Q).
9. Remove the brain, ventral nerve cord (VNC), trachea, and gut using a Dumont #5 forceps (Figure 4J,K,Q). Be careful not to disturb the developing IFMs, which are directly underneath the trachea and VNC attached to the epidermis on the dorsal portion of the thorax.
10. Clip the tip of a 200 µL pipette tip to create a wider opening. Use a 200 µL pipette with the clipped tip to gently wash 1x PBS buffer over the thorax to remove most of the remaining fat bodies to expose the IFMs (Figure 4L,M). Be gentle to avoid tearing the tendon attachments that connect the myofibers to the epidermis.
11. Use a Vannas spring scissors to cut down the midline of the dorsal thorax, creating two "leaflets" containing IFMs (Figure 4N).
12. Cut the leaflets of the dorsal thorax away from the abdomen using a scissors (Figure 4O, Q), and transfer the leaflets to 300 µL of fixative solution in a black glass dish using a Dumont #5 forceps (Figure 4P). Grasp the leaflets at the head case with the forceps to avoid damaging the IFM myofibers or tendons.
13. Fix for the desired length of time (typically 15 min). Use a 200 µL pipette to carefully remove fixative and wash in 500 µL of 0.05% PBS-T. Proceed with blocking and immunostaining (see step 4 below).
    NOTE: Do not nutate or rock samples in black glass dishes. Muscle-tendon attachments at early pupal stages are very fragile and will tear with constant buffer agitation, resulting in loss of IFMs. Slowly and carefully pipette around the edges of the buffer in the dish to avoid sucking samples into the pipette tip when changing solutions.

4. Immunostaining and histochemistry of fixed IFM samples

1. Assemble the necessary supplies, including a pipette, pipette tips, and required antibodies or stains. Prepare 0.05% PBS-T, blocking solution, and antibody/stain mixtures (see Supplementary File 1).
2. Pipette the wash solution (0.05% PBS-T) off the samples and add the blocking solution. Incubate samples in blocking solution for at least 30 min at room temperature (with rocking for steps 1-2, without rocking for step 3).
   NOTE: Samples can alternatively be blocked overnight at 4 °C. In this protocol, samples were blocked in 5% normal goat serum (NGS) in 0.05% PBS-T. Further information on blocking solution options is provided in Supplementary File 1.
3. Remove blocking solution from the thoraces using a pipette. Add primary antibody mixture and incubate at least 2 h at room temperature or overnight at 4 °C (with rocking for steps 1-2, without rocking for step 3).
   NOTE: This protocol is compatible with the majority of primary antibodies. Antibodies should be diluted in blocking solution. Primary antibody dilutions tested with this protocol range from 1:10 to 1:2000 and are determined experimentally for each individual antibody. Further details on the optimization of primary antibody staining are provided in Supplementary File 1.
4. Remove the primary antibody mixture using a pipette and wash the samples 3x for 10 min each, in 0.05% PBS-T.
5. Add secondary antibody or stain mixture to the sample. Cover the sample with a piece of aluminum foil to minimize bleaching. Incubate at least 2 h at room temperature or overnight at 4 °C (with rocking for step 1-2, without rocking for step 3).
   NOTE: Most secondary antibodies and fluorescent cellular stains can be used with this protocol. Many secondary antibodies are used at a 1:500 dilution. In the representative data below (see Representative Results), nuclei were labeled with DAPI (1 mg/mL stock, diluted 1:1000 in 0.05% PBS-T) and F-actin with rhodamine phalloidin (300 units/mL stock, diluted 1:500 in 0.05% PBS-T).
6. Remove the secondary/stain mixture using a pipette and wash the samples 4x for 10 min each, in 0.05% PBS-T at room temperature. Proceed to mounting (step 5) directly after washing.

5. Mounting stained samples on slides for microscopy analysis (Figure 5)

1. Assemble the necessary supplies including a paint brush, forceps, frosted microscope slides, coverslips (#1 thickness), clear nail polish, mounting medium, and lint-free wipes (see Table of Materials).
2. Label the frosted writing area of a microscope slide with the relevant sample number or preferred identifying information (date, genotype, antibody, slide number, etc.).
3. Add coverslip spacers (#1 coverslips), leaving about 1 cm between the coverslips for sample placement (Figure 5A-C). Use a small drop of 50% glycerol to hold the spacers in place (Figure 5A, D).
   NOTE: Spacers prevent sample distortion when mounting thick samples. Adult thoraces typically require a 2-coverslip thick spacer, 48 h APF thoraces a 1-coverslip thick spacer, and open-book leaflets no spacer.
4. Add a drop of mounting medium to the well between the coverslip spacers, or to the center of the microscope slide if mounting early pupal samples (Figure 5C-D). Place the slide under a stereo dissecting microscope.
   NOTE: Multiple types of mounting media are available. If using a self-hardening media, consider one with a cure-time of several hours to allow sufficient time to properly orient samples before the media hardens. Mounting media have been shown to influence sarcomere dimensions^43,44,45, so the same reagent should be used for all samples in an experiment.
5. Use a pipette to remove as much wash buffer from the samples as possible. Immediately transfer the samples using a forceps or paintbrush to the drop of mounting medium on the slide (Figure 5E).
6. Using Dumont #5 forceps, organize the samples into rows and columns, ensure the samples are non-overlapping, and orient the thoraces with the IFMs facing up (Figure 5F,J,Q,S). Orientation may require carefully flipping over samples where the bristles/cuticle are facing up (Figure 5G,H,K, L,R). Be careful not to touch the IFMs, as contact will damage sarcomere structure (see Representative Results).
7. Carefully add a #1 coverslip (18 x 18 mm or 22 x 22 mm) over the thorax samples (Figure 5M). If using spacers, gently press the coverslip until it contacts the spacers on either side of the samples (Figure 5N).
8. Use a pipette tip or dropper to back-fill the sample area with mounting medium until all thoraces are encased (Figure 5O). Avoid creating air bubbles that will disrupt image acquisition.
9. Use clear nail polish to seal the wet-mount. Remember to seal around both the sample coverslip as well as the spacer coverslips (Figure 5P,S).
   NOTE: Avoid using top coat or quick-dry nail polish, as they seal poorly and air bubbles will form as the mounting medium evaporates.
10. Dry slides for 10 min at room temperature. Store in a slide case at 4 °C. Slides can be imaged for several days or weeks.
11. Image samples on a fluorescent microscope, confocal microscope, or other system according to the experimental design. Protocols for acquisition and image analysis are available elsewhere^{14,30,46,47,48}.

The dissection protocol presented above can be used to generate high-quality samples for microscopy analysis of IFM myofiber and sarcomere morphology from as early as 8 h after puparium formation (APF) using the open-book method (step 3) through adult stages using the hemithorax dissection approach (steps 1 and 2). These dissections have been applied to investigate muscle and sarcomere formation18,42,49, transcription factor function50,51, and RNA regulation37,38, among others. The representative results provided below demonstrate the compatibility of this protocol with different fixation methods, detail common dissection artifacts, and use the SmnE33 mutant phenotype to illustrate the utility of this protocol to address morphological and cell biological questions in the IFM model.

IFM dissection is compatible with multiple fixation methods

A critical aspect of sample preparation for histochemistry or immunofluorescence samples is accurate preservation of cellular structures through fixation. Fixatives preserve tissue structure and morphology by preventing degradation and stabilizing protein and lipid structures through crosslinking52,53. Different classes of fixatives, for example, aldehyde-based versus organic solvents, have different penetration and cross-linking efficiencies and differentially impact antibody-target recognition by targeting distinct functional groups53,54. IFM dissections are compatible with multiple fixation methods, including 4% paraformaldehyde (PFA), 9% glyoxal, and methanol fixation (Figure 6), emphasizing the general utility of this dissection protocol.

To compare sarcomere size and morphology with the application of different fixation methods, hemithorax dissections of 1 day adult w1118 flies were performed. 4% PFA fixation in either phosphate buffered saline (PBS) (Figure 6A,E) or relaxing solution (RS) (Figure 6B,F) was compared. Relaxing solution contains ATP and is recommended for use when quantifying sarcomere dimensions to preserve sarcomeres in their relaxed state20,55 (see Supplementary File 1). 4% PFA fixation effectively preserves sarcomere morphology, but a significant difference in sarcomere width was seen between 4% PFA fixation in PBS vs. RS (Figure 6I,J). Fixation in methanol preserved myofiber and sarcomere structure (Figure 6C,G), but sarcomeres were significantly shorter and thinner than with 4% PFA fixation (Figure 6I,J). IFMs could be effectively preserved with an overnight 9% glyoxal fixation (Figure 6D,H) (3% glyoxal fixation was ineffective), which also significantly altered sarcomere width as compared to 4% PFA fixation (Figure 6I,J). In conclusion, IFM hemithoraces can be effectively preserved with multiple fixatives. However, as both fixative and buffer can influence the measurement of sarcomere dimensions, experiments should be designed with proper controls and a well-defined and consistent fixation protocol.

Common dissection and fixation artifacts in IFMs

One challenge with microscopy, especially for new users and trainees, is distinguishing technical artifacts from bone-fide phenotypes. To aid with this distinction in Drosophila IFMs, representative data of the most commonly observed artifacts and dissection failures is presented (Figure 6). Fixation protocols require optimization of the type, concentration, and length of fixation to avoid under- or over-fixation52,53,54. Using a fixation time course of w1118 1 day adult hemithorax dissections incubated in 4% PFA in relaxing solution, it was observed that while 15- or 30-min fixation time accurately preserves sarcomere morphology, a 7-min fixation produces inconsistent sarcomere morphologies (Figure 6K-M). In addition, the tissue is less rigid with a shorter fixation time, making it more difficult to cut and bisect the thorax without damaging the IFMs. Additional artifacts can arise during both the cut and the mounting process. IFM myofiber structure can appear irregular if the blade is dull and causes ripping or fraying (Figure 6N) or if the blade slips or the cut is angled (Figure 6O). If forceps come into direct contact with fixed IFMs during handling or mounting, they make an impression that can lead to divots or stretching and can pull the sarcomeres apart (Figure 6P). Stretching or deformation of the thorax, either during cutting, handling, or mounting, also pull the sarcomeres apart and result in loss of a clearly demarcated Z-disc (Figure 6Q). Lastly, "sawing" or "rubbing" of the IFMs with a forceps or blade leads to abrasion and unraveling, fraying, and disorganization of myofibrils, especially on the surface of the myofiber (Figure 6R). Potential users should become familiar with these technical artifacts and exclude them from phenotypic analysis.

Developmental progression of the SmnE33 phenotype in IFMs

Open-book and hemithorax dissections are particularly useful to track the developmental trajectory of a muscle phenotype and to pinpoint when in development a gene of interest acts or myofibril phenotypes arise. The IFM developmental phenotype of SmnE33, a hypomorphic allele of the RNA-binding protein Survival motor neuron (Smn)39 is presented as representative data illustrating the broad utility of open-book and hemithorax dissections.

The small nuclear ribonucleoproteins (snRNPs) that form the spliceosome, the macromolecular complex which catalyzes the splicing reaction to generate mRNAs7, are assembled by the SMN complex. The SMN complex, consisting of Survival motor neuron (Smn) and Gemin proteins, is required for viability in all organisms56. Reduced SMN levels in humans lead to the severe inherited neuromuscular disorder Spinal Motor Atrophy (SMA)21. Drosophila has been used as a model to understand the etiology of SMA and cellular function of Smn, including the development of the hypomorphic allele SmnE33. SmnE33 flies are viable and fertile, but cannot fly or jump39. While SmnE33 flies have motor neuron arborization defects, they also display disorganized IFM structure and a loss of flight-muscle specific Actin88F expression39,57. SMN is co-localized with alpha-actinin at the Z-disc in flies and mice39, suggesting that Smn may have a muscle-specific function.

To gain a better understanding of when myofibril structure is lost and Smn function is required in developing muscle, a developmental time course analysis of the SmnE33 phenotype in IFMs at 26 h APF, 72 h APF, and 1 to 5-day adult was performed. While control w1118 IFMs at both 72 h APF and in adult have strong phalloidin stained F-actin signal (Figure 7E,E',I,I') and regular sarcomere structure (Figure 7G,G',K,K'), phalloidin signal is dramatically reduced (Figure 7F,F',J,J') and sarcomere structures are absent (Figure 7H,H',L,L') in SmnE33 IFMs. F-actin is instead observed in net-like structures around nuclei or in irregular bundles or "starburst" structures in the cytoplasm (Figure 7H,H',L,L'), consistent with a loss of Act88F expression18,39 and an inability to assemble thin-filaments. Open-book dissections were performed to examine the SmnE33 phenotype at 26 h APF and revealed that early IFM development proceeds normally in SmnE33. Comparable to w1118 control IFMs (Figure 7A,A',C,C'), SmnE33 flies have 6 dorsal longitudinal IFM fibers per hemithorax, form organized F-actin cables at the myofiber periphery prior to myofibrillogenesis (Figure 7B,B'), and display a mesh-like F-actin network throughout the myofiber (Figure 7D,D'). These results indicate that initial stages of IFM differentiation proceed normally, while Smn is necessary in later stages of IFM development to sustain Act88F expression and build sarcomeres. Importantly, these representative results illustrate the phenotypic detail and resolution that can be achieved by open-book and hemithorax dissections.

Figure 1: Hemi-section dissection of adult IFMs. (A) Position an adult fly in 1x PBS on a microscope slide. (B) Using a forceps (cyan, dot) to stabilize the fly, remove the head with a scissors (orange). (C-E) Remove the left (C) and right (D) wings, and the abdomen (E). (F) Discard the head, abdomen, and wings, and transfer the thorax to fixative. (G,H) Orient a fixed thorax on a slide in a drop of 1x PBS-T with the posterior (P) and the scutellum pointing up (G), using a forceps to stabilize the orientation (H). (I-K) Use a cryostat blade to cut the thorax in half (I-J), to generate two thorax hemi-sections (K). (L) Diagram of a cryostat blade and a thorax, illustrating the position of the sagittal cut to produce hemi-sections. The blade is slid to the right to notch the scutellum of the thorax, and then moved downward in a smooth motion (yellow arrows) to cut the thorax (dotted red line). (M,N) Schematic of a coronal (M) and transverse (N) view of the thorax, illustrating the position of the major muscle groups and the position of the cut (between the yellow triangles). The orientation of the thorax is indicated (L, left; R, right; V, ventral; D, dorsal; A, anterior; P, posterior). Scale bars = 1 mm. Please click here to view a larger version of this figure.

Figure 2: Pupal staging and features of pupal development. (A) Male and (B) female pupae can be distinguished as white pre-pupae at 0-2 h APF. Male pupae have paired, round, translucent structures (the developing testis) towards the posterior (arrows). (C) Table of features that serve as hallmarks of pupal development. Features include maturation of the pupal case, head eversion, and detachment of the pupa from the pupal case, development of eye pigmentation, pigmentation of the bristles, wings, legs, epandrium, and cuticle, and visibility of the virgin spot (meconium). Eye color and pigmentation darken progressively. The timing and stages of pupal development labeled at the top of each column in the table (timepoints denoted in black text, corresponding stage in pupal development below in blue text) were previously defined by Bainbridge and Bownes19,41. (D-E) Time course of pupal development at 0-2 h, 24 h, 48 h, 72 h, and 96 h APF in an intact pupa with red eyes (D; Mef2-Gal4, fln-GFP) and in a pupa with orange eyes dissected out of the pupal case (E; Fln-Gal4). Note the progressive darkening of the eyes and bristles from 48 h to 96 h APF. (F) Examples of pupal lethality at early, middle, and late timepoints of development. Pharate lethal flies are fully formed, but fail to completely eclose from the pupal case. Scale bars = 1 mm. Please click here to view a larger version of this figure.

Figure 3: Hemi-section dissection of pupal IFMs (>48 h APF). (A-H) Removal of a pupa from the pupal case. Affix pupae to a strip of double-stick tape (A,B). Tease open at the anterior ridge (C) and remove the operculum (D) using a pair of Dumont #5 forceps (blue, arrow denotes direction of movement, dot is stationary). Use forceps to cut open (E) and peel away (F) the pupal case in strips (G). Strips of the pupal case are adhered to the double-stick tape. After exposing the abdomen, lift the pupa out of the case (H). (I) Schematic illustrating the process of dissecting a pupa out of the pupal case. The figure panels corresponding to each step in the schematic are labeled (bottom). The red dotted line marks where the forceps can be inserted without damaging the pupa to cut open the pupal case. (J-L) After removing the pupa from the case (J) and transferring to a microscope slide in a drop of 1x PBS (K), use a scissors to remove the abdomen (L). (M) Discard the abdomen, and transfer the thorax to fixative. (N) After fixation, transfer pupal thoraces to a slide in a drop of 1x PBS-T. Orient the pupa dorsal side up and stabilize with a forceps. (O-P) Use a cryostat blade (O) to cut a pupal hemi-section (P). Placement of the cut is the same as Figure 1 M-N. Scale bars = 1 mm. Please click here to view a larger version of this figure.

Figure 4: Open-book dissection of pupal IFMs (<48 h APF). (A) After removing the pupa from the pupal case as shown in Figure 3 A-I, transfer the pupa to 1x PBS in a black silicon dissecting dish. (B-D) Gently push the pupa down to the surface of the silicon dish using a forceps (B), and pin ventral side up using two insect pins (C,D). (E) Open the basal membrane (bm) and cuticle of the head using a scissors (orange). (F,G) Cut along the right (F) and left (G) side. The position of the cut is diagrammed in (Q). (H,I) Lift the ventral section with a forceps (H) and remove with a scissors (I). (J,K) Use a forceps to remove the brain (J), the lateral trunk trachea, and gut (K). (L-M) Use a gentle stream of buffer from a pipette to remove fat bodies and expose the IFMs. (N) Cut the thorax into two leaflets. (O,P) Cut off the leaflets and transfer them to fixative. (Q) Schematic summarizing the steps in an open-book dissection. The figure panels corresponding to each step are labeled (bottom). The top and side views are provided to illustrate placement of the cuts (dotted red line) on the left and right side of the pupa. The wings, legs, and proboscis (labeled) are used to distinguish the dorsal and ventral sides of the pupa. Please click here to view a larger version of this figure.

Figure 5: Sample mounting on microscope slides. (A-D) Coverslip spacers are used to mount thick thorax hemi-section samples. Glycerol (A) is used to affix spacers (#1 coverslips) to a labeled slide (B), and samples are mounted in mounting medium in the space between the spacers (C). Late pupae and adult thoraces require two #1 coverslip spacers (D). Mid-pupal timepoints require a single #1 coverslip spacer, and early pupal dissections can be mounted with no spacer. (E-H) Adult hemithorax samples are transferred into mounting medium with a forceps or paintbrush (E). Thoraces are initially randomly oriented (F), and may need to be flipped using a forceps (G) so that the IFMs are oriented up towards the coverslip (H). (I-L) Leaflets from early-pupal open book dissections are transferred for mounting using a forceps (I). Leaflets are randomly oriented (J) and can be flipped using a forceps (K) so that IFMs are facing up towards the coverslip (L). (M-P) After samples are properly oriented, place a coverslip over the samples (M) and tap it even against the spacers (N). Fill around the samples with mounting medium (O), being careful to avoid forming bubbles. Seal all open edges of the coverslip and spacers with nail polish (P) to avoid evaporation of the mounting medium. (Q-R) Images of properly mounted adult thorax samples (Q, 10x magnification) oriented with the IFMs facing up towards the coverslip (R, 20x magnification). (S) A schematic of the completed slide, with samples oriented IFM-up between spacers and nail polish sealing all open edges. Please click here to view a larger version of this figure.

Figure 6: IFM dissections are compatible with different fixatives. (A-H) Confocal z-projection of myofiber structure (A-D) or single-plane images of myofibril and sarcomere structure (E-H) of control w1118 adult IFM. Hemithorax dissection is compatible with multiple fixation methods, including 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) (A,E) or relaxing solution (RS) (B,F), methanol (C,G), or 9% glyoxal solution (D,H). DAPI (1:1000), blue; phalloidin (1:500) stained F-actin, grey. (I,J) Quantification of sarcomere length (I) and width (J) from E-H. The fixation method and buffer can significantly impact measurement of sarcomere length and width. A sarcomere length of 3.040 ± 0.2935 µm with methanol fixation was significantly shorter than measured lengths of 3.271 ± 0.2736 µm, 3.190 ± 0.2586 µm, and 3.217 ± 0.2023 µm with PFA (PBS), PFA (RS) and 9% glyoxal fixation, respectively (p < 0.001). Sarcomere width was significantly different between all fixation methods tested (PFA (PBS), 1.410 ± 0.1331 µm; PFA (RS), 1.284 ± 0.2514 µm; methanol, 1.280 ± 0.1538 µm; and 9% glyoxal fixation, 1.137 ± 0.2032 µm). Boxplots are shown with Tukey whiskers with outlier data points marked as black dots. Significance was determined by ANOVA with a post hoc Tukey test (**, p < 0.01; ***, p < 0.001). (K-M) Single-plane confocal images of a fixation time course of adult w1118 in 4% PFA in RS fixation, demonstrating that fixation times of 15 (L) or 30 (M) min, as compared to 7 min (K), result in well-preserved and consistent sarcomere structure. (N,O) Z-stack projections of adult Act88F-Gal4 IFMs fixed in 4% PFA demonstrating dissection artifacts in myofibers.Common artifacts include cut, frayed, or partial myofibers from a dull blade (N) or an angled cut (O). (P-R) Single-plane confocal images of adult w1118 samples fixed in 4% PFA in PBS demonstrating common dissection artifacts in sarcomeres and myofibrils. Common technical artifacts include irregular stretching of sarcomeres due to contact with forceps while mounting (P), pulling Z-discs out of register by stretching myofibers during longitudinal cuts (Q), and frayed, disorganized, or curling myofibrils due to abrasion or a dull blade (R). DAPI (1:1000), blue; phalloidin (1:500) stained F-actin, grey. Scale bar = 100 µm (A-D, N-O), 5 µm (E-H, K-M, P-R). Please click here to view a larger version of this figure.

Figure 7: Application of dissections to investigate the developmental IFM phenotype of SmnE33. (A-D) Muscle structure in early pupae at 26 h APF. Single-plane confocal image of myofiber structure (A, A', B, B') and myofibril and sarcomere structure (C, C', D, D') in control (A, A', C, C') and SmnE33(B, B', D, D'). Both control and SmnE33 have six IFM myofibers per hemithorax and form F-actin cables. (E-H) Muscle structure in late pupae at 72 h APF. Z-projection image of myofiber structure in control (E, E') and SmnE33 (F, F'). IFMs in SmnE33 are present based on DAPI staining but lack a strong F-actin signal. Single-plane confocal images reveal that control IFMs have a highly organized sarcomere structure (G, G'). By contrast, SmnE33 IFMs (H, H') have abnormal star-like F-actin structures (yellow arrows) and lack the organized sarcomere structure observed in control IFMs. (I-L) Adult muscle structure in control (I, I', K, K') and SmnE33(J, J', L, L'). Z-projection (I,J) and single-plane confocal images (K,L) reveal greatly reduced F-actin content and abnormal actin structures (yellow arrows) in SmnE33IFMs. DAPI (1:1000), blue; phalloidin (1:500) stained F-actin, magenta or grey. Scale bar = 50 µm (A, B), 10 µm (C, D, G, H, K, L), 100 µm (E, F, I, J). Please click here to view a larger version of this figure.

Supplementary File 1: A detailed description of the fixation and staining methods used in the text and in particular to generate the data shown in Figure 6 and Figure 7. Additional information on determining dilutions of primary antibodies is included. A list of buffer components and recipes used in this protocol is also provided. These data motivate the dissection protocol and demonstrate its utility for confocal microscopy and analysis of developmental IFM phenotypes. Please click here to download this file.

Supplementary Figure 1: Flow diagram of steps in hemithorax dissection of adult IFMs. Textual summary of the steps in Figure 1 to prepare hemithorax IFM sections. After dissection, fixation, and thorax bisection, sections are stained for microscopy analysis. Please click here to download this figure.

Supplementary Figure 2: Flow diagram of steps in hemithorax dissection of pupal IFMs. Textual summary of the steps in Figure 3 to prepare hemithorax sections of pupal IFMs. After removal from the pupal case, pupa are fixed, bisected, and stained for microscopy. Please click here to download this figure.

Supplementary Figure 3: Flow diagram of steps in open book dissection of early pupal IFMs. Textual summary of the steps shown in Figure 4 to perform open-book dissection of pupal IFMs before 48 h APF. After dissection and fixation, epithelial leaflets with attached IFMs are stained for microscopy. Please click here to download this figure.

This protocol presents a procedure for hemithorax dissection of D. melanogaster adult (Figure 1) and late pupal (48 h - 96 h APF) (Figure 3) IFMs as well as open-book dissection of IFMs at early pupal (8 h - 48 h APF) (Figure 4) timepoints. This protocol is optimal for immunohistochemistry and fluorescence microscopy applications. The protocol is illustrated with schematic diagrams and stepwise photographs of the dissection procedure. Overview diagrams further make the protocol accessible to scientists new to the Drosophila field as well as undergraduate researchers and early-stage trainees. Representative examples of poor fixation and sample preparation artifacts are included to enable protocol users to distinguish artifacts from bona fide sarcomere phenotypes.

The most difficult step in IFM hemithorax dissection is bisection of the thorax (Figure 1I,J,L and Figure 3O). This step requires time and practice to master. For additional stability while cutting, individual thoraces can be placed on double-stick tape and cut³⁶, but care should be taken to avoid desiccation of the IFMs. It is also possible to cut from the ventral side of the thorax, with the dorsal side oriented down, so new users can test which approach is most effective with their setup. Cuts should be performed with new, sharp blades to avoid damaging the IFMs, and a blade should be replaced when it becomes dull. Microtome (cryostat) blades are used to cleanly cut the thorax in this protocol, as compared to other protocols that employ a needle, forceps, or a scalpel blade^24,32,36. Razor blades are another alternative to cleanly bisect the thorax. Fixation is also important prior to bisection, as lightly fixed tissue, although stably preserved, may be "squishy" and more difficult to cut. Fixation times of 30-60 min balance antigen availability and tissue stiffness facilitating cuts, and can be shortened to 15 min with experience.

One limitation of hemithorax preparations is antibody penetration. Hemithoraces are thick samples³², and the best quality confocal micrographs are obtained from the top 10-20 µm of the IFMs^42,58. Too low a detergent concentration or too short a permeabilization time can lead to weak and inconsistent staining. Antibody penetration can also be improved by a longer (2-3 d) incubation in primary antibody. A two-photon system or an objective with a larger working distance can also increase imaging depth⁵⁹. If the experimenter needs to image the full diameter of the IFM myofiber, an alternate approach such as microtome^24,58 or cryosectioning²⁷ in the sagittal plane should be considered.

An important aspect that is not addressed in the main protocol is image data analysis. For analysis of IFM myofibers in 10x or 20x images (Figure 6A-D, Figure 7A,B,E,F,I,J), standard analysis includes determining if the myofibers are attached and if the gross morphology of the myofiber is intact^37,38. Examples of parameters that can be quantified, depending on the experiment, include the length, width, number, number attached, or number of ripped myofibers^37,60. A major advantage of this protocol is compatibility with fluorescence microscopy and the ability to analyze sarcomere and myofibril morphology at higher magnifications of 60x or 100x (Figure 6E-H, Figure 7C,D,G,H, K,L). Sarcomere morphology should be analyzed in samples free of technical artifacts (Figure 6K-P). A number of categories of possible myofibril and sarcomere phenotypes has been previously defined⁶⁰. Most commonly, sarcomere length and width are measured either by drawing and measuring a line from Z-disc to Z-disc in image processing software, or alternately using one of several automated tools that are freely available^18,45,61. Assessment of sarcomere morphology can be enhanced by including markers that label the Z-disc or M-line, or the thick or thin filaments^30,34,58. These data can be used to quantitatively describe myofiber, myofibril, and sarcomere phenotypes between a control and test sample prepared using this dissection protocol.

Protocol users should be aware that while fixation preserves myofiber and sarcomere morphology, fixative as well as buffers and the mounting medium may cause tissue shrinkage or swelling and can influence the measurement of morphological parameters⁴⁵. The glutaraldehyde fixation and dehydration necessary for electron microscopy is known to shrink sarcomere diameter⁴³, while fiber skinning and glyceration have been shown to increase sarcomere diameter^44,45. This phenomenon may underlie the differences in sarcomere parameters in the literature and emphasizes the importance of including control and test samples in each experiment. Control and test IFMs should be processed simultaneously in the same buffers for accurate comparison of sarcomere quantitative parameters.

This detailed protocol aims to make adult and pupal IFM dissection for immunohistochemistry and fluorescent microscopy more accessible to beginning and advanced Drosophilists. This protocol can be adapted for use with other insects as well as for other imaging techniques such as cryo-electron microscopy (cryo-EM), single-molecule fluorescence in situ hybridization (smFISH), or spatial transcriptomics. The combination of powerful Drosophila genetics tools with fluorescence microscopy offers unique opportunities to investigate conserved principles of myogenesis and muscle function. As IFMs are also amenable to molecular and biochemical approaches^26,27, future studies linking molecular phenotypes to muscle morphology, cell biology, and function will provide a deeper understanding of myogenesis and the etiology of muscle disorders.