Name: Dissection of Drosophila melanogaster Indirect Flight Muscles for Microscopy Approaches (Video) | JoVE Journal
Uploaded: 2025-11-07T14:20:00.000+00:00
Duration: 9 min 47 s
Description: Learn dissection techniques for Drosophila melanogaster indirect flight muscles to study sarcomere formation and RNA regulation in myogenesis. This protocol enables advanced microscopy analysis of muscle development.

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The authors thank Gregory Matera for providing the SmnE33 allele and for helpful discussions on Smn. The authors thank Frank Schnorrer, Cornelia Schoenbauer, Manuela Weitkunat, and Aynur Kaya-Copur for helpful discussions and support. The authors acknowledge the Bloomington stock centre for providing flies. This work was supported by start-up funding from the University of Missouri Kansas City (UMKC) School of Science and Engineering, Division of Biological and Biomedical Systems (MLS), the UMKC Funding for Excellence Program (MLS), and the UMKC Office of Undergraduate Research and Creative Scholarship (SF, MLS).

The authors have nothing to disclose.

This protocol presents a procedure for hemithorax dissection of D. melanogaster adult (Figure 1)&#160;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 blade24,32,36. 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 &#34;squishy&#34; 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 &#181;m of the IFMs42,58. 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 microtome24,58 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 intact37,38. Examples of parameters that can be quantified, depending on the experiment, include the length, width, number, number attached, or number of ripped myofibers37,60. 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 available18,45,61. Assessment of sarcomere morphology can be enhanced by including markers that label the Z-disc or M-line, or the thick or thin filaments30,34,58. 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 diameter44,45. 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 approaches26,27, 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.

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 mechanotransduction5,6,7 that instruct and fine-tune muscle development and function. Many of these cellular processes are disrupted in human myopathies and neuromuscular disorders8,9, 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 proteins10,11 coupled to technological advances in image acquisition and resolution12,13 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 &#176;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 &#176;C, and at approximately 100 h APF (females at about 98 h APF and males at about 102 h APF) at 25 &#176;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 approaches8,20. 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 insights21,22. 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 metals3,23,24,25, but are not optimal for light microscopy. Protocols developed to rapidly dissect IFMs out of the thorax for use in biochemical assays or molecular analyses26,27 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 muscle20,28, 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 process29,30, 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 staining27,31,32, 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 microscopy20,33,34. Other examples include protocols to dissect intact IFM fibers from the thorax32,35 or to generate whole-mount or hemithorax preparations20,32,36 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 SmnE33, a hypomorphic allele of survival motor neuron (Smn) used as a fly model of&#160;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.

<table><tbody><tr><td>60 mm tissue culture dishes</td><td>Fisher Scientific</td><td>FB012921</td><td>60 mm, polystyrene</td></tr><tr><td>Acetic Acid, Glacial (Certified ACS)</td><td>Fisher Scientific</td><td>A38S-212</td><td></td></tr><tr><td>Adenosine Triphosphate (ATP)</td><td>Millipore Sigma</td><td>A1852</td><td></td></tr><tr><td>Aluminum foil, heavy duty</td><td>Amazon</td><td></td><td>12 in. x 1000 ft.</td></tr><tr><td>Black silicon dissecting dishes: activated charcoal powder</td><td>Millipore Sigma</td><td>C9157</td><td>Also available from most pharmacies</td></tr><tr><td>Black silicon dissecting dishes: Sylgard 184</td><td>Millipore Sigma</td><td>761036</td><td>To make silicon dissecting dishes, combine Sylgard components in a ziploc bag and mix well. Add activated charcoal powder (~200 mg) to Sylgard (~50 g), mix well, cut the corner of the bag, fill 60 mm culture dishes, remove bubbles with a pipette tip, and cure overnight.&amp;nbsp;</td></tr><tr><td>Cardboard slide tray, Research Products International Corp</td><td>Fisher Scientific</td><td>50-136-7558</td><td></td></tr><tr><td>Confocal microscope, Nikon AXR inverted confocal</td><td>Nikon</td><td></td><td>&lt;a href=&quot;http://www.microscope.healthcare.nikon.com/&quot;&gt;www.microscope.healthcare.nikon.com/&lt;/a&gt;</td></tr><tr><td>Confocal microscope, Zeiss LSM 780 inverted confocal</td><td>Zeiss</td><td></td><td>&lt;a href=&quot;http://www.zeiss.com/&quot;&gt;www.zeiss.com&lt;/a&gt;</td></tr><tr><td>Darwin Insect Rearing Chamber (Incubator)</td><td>Fisher Scientific/Darwin Chambers</td><td>IN034</td><td>light, temperature, and humidity controlled incubator</td></tr><tr><td>dissecting dish (black glass)</td><td>Mikroskop Technik Diethelm</td><td>398011</td><td>Lymphbecken (dyeing bowls/lymph tanks), black glass, 4x4 cm, with clear glass cover</td></tr><tr><td>Dumont #3 Forceps</td><td>Fine Science Tools</td><td>11231-30</td><td>Dumoxel straight tip 12 cm forceps with 0.17 x 0.1 mm tip&amp;nbsp;</td></tr><tr><td>Dumont #5 Forceps</td><td>Fine Science Tools</td><td>11252-20</td><td>Inox straight tip 11 cm forceps, Biology grade with 0.05 x 0.02 mm tip&amp;nbsp;</td></tr><tr><td>EGTA (Egtazic Acid)</td><td>Millipore Sigma</td><td>324626</td><td></td></tr><tr><td>Ethanol, Absolute (200 Proof)</td><td>Fisher Scientific</td><td>BP28184</td><td>Molecular Biology Grade</td></tr><tr><td>Fisherbrand Premium Cover Glass (18 x 18 mm), coverslips</td><td>Fisher Scientific</td><td>12548A</td><td>Glass, Square, #1 thickness (0.13 - 0.16 mm)</td></tr><tr><td>Fisherbrand Premium Cover Glass (22 x 22 mm), coverslips</td><td>Fisher Scientific</td><td>12548B</td><td>Glass, Square, #1 thickness (0.13 - 0.16 mm)</td></tr><tr><td>Flat bottom cell culture plates (24-well)</td><td>Fisher Scientific</td><td>7200740</td><td></td></tr><tr><td>fluorescent dissecting microscope camera, Infinity8 camera</td><td>Teledyne (Visual Dynamix)</td><td></td><td>6 megapixel, low-light, monochromatic</td></tr><tr><td>Fly: Smn[E33]</td><td></td><td></td><td>Fly stock; gift of Gregory Matera</td></tr><tr><td>Fly: w[1118]</td><td>Bloomington Stock Center</td><td>RRID:BDSC_3605</td><td>Fly stock</td></tr><tr><td>glycerol</td><td>Fisher Scientific</td><td>BP229-1</td><td>Molecular Biology Grade</td></tr><tr><td>Glyoxal solution (40 wt. % in H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Millipore Sigma</td><td>128465</td><td>&amp;nbsp;This is 40% stock solution</td></tr><tr><td>Image processing software, Affinity Designer 2</td><td>Affinity</td><td></td><td>https://affinity.serif.com/en-us/</td></tr><tr><td>Image processing software, Image J (Fiji)</td><td>ImageJ2</td><td></td><td>https://imagej.net/software/fiji/</td></tr><tr><td>Insect pins</td><td>Fine Science Tools</td><td>26002-10</td><td>stainless steel, 0.1 mm diameter (minutien pins)</td></tr><tr><td>Magnesium Chloride (MgCl2)</td><td>Fisher Scientific</td><td>AA12315A1</td><td></td></tr><tr><td>Methanol (HPLC)</td><td>Fisher Chemical (Fisher Scientific)</td><td>A452-4</td><td>Chill at -20 ? before use</td></tr><tr><td>Microscope slides, frosted</td><td>Fisher Scientific</td><td>12-550-400</td><td>writing area, uncharged, 75 mm x 25 mm x 1 mm</td></tr><tr><td>Microscope slides, plain glass</td><td>Fisher Scientific</td><td>12-544-4</td><td>Precleaned, 75 mm x 25 mm</td></tr><tr><td>MX35 Ultra microtome blades</td><td>epredia (Fisher Scientific)</td><td>3053835</td><td>Low profile, 34&amp;deg; cutting angle; C35 feather 80mm blades work well</td></tr><tr><td>Nail polish, clear (Sally Hansen Xtreme Wear Nail Polish)</td><td>Amazon</td><td></td><td>0.4 fl. oz.</td></tr><tr><td>Normal goat serum</td><td>Millipore Sigma</td><td>S26-LITER</td><td></td></tr><tr><td>Nutator (Mini Nutating Rocker, Benchmark)</td><td>Midwest Scientific</td><td>H3D1020</td><td>24 rpm, fixed; 8 x 6 in. platform with dimpled mat</td></tr><tr><td>Paintbrush (Round No. 0)</td><td>Amazon</td><td></td><td>Round No. 0 or similar brush from any art supply</td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Millipore Sigma</td><td>158127-500G</td><td></td></tr><tr><td>Phosphate Buffered Saline (PBS): KCl</td><td>Millipore Sigma</td><td>529552-250GM</td><td>Prepare PBS according to Cold Spring Harb Protoc; 2006; doi:10.1101/pdb.rec8247</td></tr><tr><td>Phosphate Buffered Saline (PBS): KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Millipore Sigma</td><td>P0662-500G</td><td>Prepare PBS according to Cold Spring Harb Protoc; 2006; doi:10.1101/pdb.rec8247</td></tr><tr><td>Phosphate Buffered Saline (PBS): Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Millipore Sigma</td><td>S9763-500G</td><td>Prepare PBS according to Cold Spring Harb Protoc; 2006; doi:10.1101/pdb.rec8247</td></tr><tr><td>Phosphate Buffered Saline (PBS): NaCl</td><td>Fisher Chemical (Fisher Scientific)</td><td>S271-1</td><td>Prepare PBS according to Cold Spring Harb Protoc; 2006; doi:10.1101/pdb.rec8247</td></tr><tr><td>PR1MA Pipette tips (10 &amp;micro;L)</td><td>Midwest Scientific</td><td>PR10XLRK-NS</td><td>Universal 10 &amp;micro;L pipette tips</td></tr><tr><td>PR1MA Pipette tips (1000 &amp;micro;L)</td><td>Midwest Scientific</td><td>PR-1000RK-FL</td><td>Universal 1000 &amp;micro;L pipette tips</td></tr><tr><td>PR1MA Pipette tips (200 &amp;micro;L)</td><td>Midwest Scientific</td><td>PR-200RK-NS</td><td>Universal 200 &amp;micro;L pipette tips</td></tr><tr><td>Rhodamine-Phalloidin</td><td>Invitrogen, Molecular Probes</td><td>R415</td><td></td></tr><tr><td>Scotch double sided tape</td><td>Scotch/3M (Staples)</td><td>649280</td><td>both sides coated with adhesive, 0.5&quot; x 7 yards, available at most office supply handlers</td></tr><tr><td>Sodium dihydrogen phosphate dihydrate (NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Fisher Scientific</td><td>AAA1131636</td><td></td></tr><tr><td>Software: GraphPad Prism</td><td>GraphPad Prism</td><td></td><td>&lt;a href=&quot;http://www.graphpad.com/&quot;&gt;www.graphpad.com&lt;/a&gt;</td></tr><tr><td>Software: Infinity Analyze 7</td><td>Teledyne (Visual Dynamix)</td><td></td><td>&lt;a href=&quot;https://www.teledynevisionsolutions.com/products/infinity-analyze/&quot;&gt;https://www.teledynevisionsolutions.com/products/infinity-analyze/&lt;/a&gt;</td></tr><tr><td>Software: Microscoft Excel</td><td>Microsoft</td><td></td><td>https://www.microsoft.com/en-us/microsoft-365/excel</td></tr><tr><td>tissue/ Kimtech Science Kimwipes Delicate Task Wipers, 1 ply</td><td>Fisher Scientific</td><td>06-666</td><td>Standard tissue wipes</td></tr><tr><td>Transfer pipette</td><td>Fisher Scientific</td><td>13-711-9AMMD</td><td>Plastic pipette</td></tr><tr><td>Trition X-100</td><td>Millipore Sigma</td><td>T9284-100ML</td><td></td></tr><tr><td>Vannas spring scissors</td><td>Fine Science Tools</td><td>15000-00</td><td>3 mm cutting edge, tip diameter 0.05 mm, length 8 cm</td></tr><tr><td>Vectashield with DAPI</td><td>Vector Laboratories</td><td>H-2000</td><td></td></tr><tr><td>Vectashield without DAPI</td><td>Vector Laboratories</td><td>H-1900</td><td></td></tr><tr><td>Whatman paper</td><td>Millipore Sigma</td><td>WHA1004070</td><td>Filter paper circles, Grade 4, 70 mm</td></tr><tr><td>z10 fluorescent stereo dissecting microscope</td><td>Visual Dynamix (Midwest Scientific)</td><td></td><td>http://www.visualdynamix.net/</td></tr><tr><td>z850 ergonomic stereo zoom dissecting microscope</td><td>Visual Dynamix (Midwest Scientific)</td><td></td><td>http://www.visualdynamix.net/</td></tr></tbody></table>

dissection of drosophila melanogaster indirect flight muscles for microscopy approaches

Dissection of Drosophila melanogaster Indirect Flight Muscles for Microscopy Approaches

The indirect flight muscles (IFMs) of Drosophila melanogaster are a powerful genetic model to explore foundational principles of myogenesis. The contractile mechanism and many sarcomere components are conserved from flies to vertebrates, enabling the study of cellular processes from transcriptional regulation and RNA processing to metabolism and mechanobiology that direct and fine-tune muscle development. Many of these cellular pathways are altered in human myopathies, and IFM studies provide relevant insight into the molecular etiology of muscle disease. In particular, flies are well-suited for microscopy to analyze myofiber and sarcomere morphology and function across the entire process of IFM myogenesis, from myoblast specification to myofibril formation, maturation, and maintenance. Here, a protocol is presented for the dissection of D. melanogaster IFMs at pupal and adult timepoints for microscopy approaches. Illustrated protocols are provided for hemithorax dissection of late pupal and adult IFMs, as well as open-book dissection of early pupal IFMs. Fixation, staining, and sample mounting procedures and common dissection artifacts are described, with representative data demonstrating the compatibility of IFM dissection with different fixation reagents. These protocols are applied to study the hypomorphic SmnE33 allele of RNA-binding protein survival motor neuron (Smn)at 26 h after puparium formation (APF), 72 h APF, and in adult IFMs, providing new insight into a Spinal Motor Atrophy (SMA) model and illustrating the general utility of this protocol. This detailed protocol facilitates access to the IFM model system and acquisition of high-quality microscopy data to investigate principles of myogenesis.

Drosophila is a powerful model to understand fundamental mechanisms of myogenesis. This protocol for the dissection and preparation of thorax hemi-sections enables microscopy analysis of indirect flight muscle from both pupal and adult stages. This protocol enables confocal imaging of cellular morphology, protein localization, muscle structure, and multiple other aspects of myogenesis.

The indirect flight muscles (IFMs) of Drosophila melanogaster are a powerful genetic model to explore foundational principles of myogenesis. The ...

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.

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Research

JoVE Journal

Biology

This protocol demonstrates how to dissect Drosophila larvae in preparation for immunohistochemistry and/or imaging of the neuromuscular junction.

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Dissection of Drosophila melanogaster Indirect Flight Muscles for Microscopy Approaches

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