Name: dissection of drosophila melanogaster flight muscles for omics approaches
Uploaded: 2019-10-17T14:00:00.000+00:00
Duration: 8 min 33 s
Description: Watch this Scientific Journal Video about Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches at JoVE.com

We are grateful to Andreas Ladurner and Frank Schnorrer for generous support. We thank Sandra Esser for excellent technical assistance and Akanksha Roy for generating the mass spectrometry data. We acknowledge the Bloomington and Vienna stock centers for providing flies. We thank the Core Facility Bioimaging for help with confocal imaging and the Zentrallabor f&#252;r Proteinanalytik for analysis of mass spectrometry samples, both at the LMU Biomedical Center (Martinsried, DE). Our work was supported by the Deutsche Forschungs Gemeinschaft (MLS, SP 1662/3-1), the Center for Integrated Protein Science Munich (CIPSM) at the Ludwig-Maximilians-University M&#252;nchen (MLS), the Frederich-Bauer Stiftung (MLS), and the International Max Planck Research School (EN).

1. Staging the Pupae
<ol>
	<li>Raise flies of the desired genotype in bottles (Figure 1E). Either make a fresh flip of the dissection stock or set a cross with at least 20 female virgin flies. Maintain bottles until the flies begin to pupate.</li>
	<li>Collect pre-pupae with a wetted paintbrush and transfer to wetted filter paper in a 60 mm Petri dish (Figure 1F).</li>
	<li>Sex the pupae, collecting the appropriate gender for the experiment (Figure 1G). Males are identified by the presence of testes, which appear as translucent balls in the otherwise opaque pupa.</li>
	<li>Label the Petri dish with the time, date, and genotype, then age the pupae to the desired stage (Figure 1H). 
	NOTE: Maintain crosses/stocks and age pupae in a temperature-controlled incubator (i.e., 25 &#176;C or 27 &#176;C for RNAi crosses, as increased Gal4 activity at higher temperatures increases knock-down efficiency40). Ensure that the humidity is sufficiently high so pupae do not dry out when aging several days.</li>
</ol>
2. IFM Dissection Before 48 h APF
<ol>
	<li>Assemble the necessary equipment including two #5 biology grade forceps, a pipette, pipette tips, dry ice, and (for RNA samples) isolation reagent (see Table of Materials). In addition, chill black dissecting dishes (see Table of Materials), 1x phosphate-buffered saline (PBS) buffer, and 1.5 mL microcentrifuge tubes on ice.</li>
	<li>Using a wetted paintbrush, transfer staged pupae to a black dissecting dish filled about two-thirds with cold 1x PBS (Figure 2A,B). Move to a fluorescent dissecting microscope. 
	NOTE: Use as many pupae as can be dissected within a 30 min time window. Depending on experience, this ranges from 3&#8211;15 pupae. See Supplemental Methods for discussion of alternatives to black dissecting dishes.</li>
	<li>Using #5 forceps, push one of the pupae to the bottom of a black dissecting dish and adjust the microscope zoom and focus to clearly see the pupa (Figure 2C).</li>
	<li>Grasp the anterior of the pupa with one forceps (Figure 2D), then poke the pupae with a single tip of the other forceps slightly off-center in the abdomen, just behind the thorax. This holds the pupa in place and prevents the IFMs from moving into the abdomen (Figure 2E). 
	NOTE: Begin timing the length of dissection from this point, as soon as pupal integrity is disrupted. Use a defined length of dissection (for example 20&#8211;30 min) to minimize muscle death and associated transcriptomic and proteomic changes. Dissect as many flies as possible in this period of time.</li>
	<li>Using the first forceps, remove the anterior half of the pupal case (Figure 2F).</li>
	<li>Use the same forceps to pinch the exposed pupae just behind the thorax, and separate the abdomen from the thorax (Figure 2G).</li>
	<li>Using the forceps, gently squeeze the anterior part of the thorax (for &#60;35 h APF) or rip open the thorax to expose the fluorescently labeled IFMs (Figure 2H). IFMs will easily detach from the epidermis, as tendon attachments at early timepoints are fragile. Discard the remaining carcass using forceps to push it to the opposite side of the dish.</li>
	<li>Repeating steps 2.3&#8211;2.7, dissect additional pupae.</li>
	<li>Collect the IFM fibers with forceps and organize them into a pile at the bottom of the black dissecting dish (Figure 2I,J). Remove any debris by pushing it out of the field of view using forceps. 
	NOTE: With practice, forceps tips can be brought into close proximity without touching each other. This technique can be used to loosely grab IFMs without destroying them. Alternate methods include gently pushing or lifting the IFMs with a single tip or completely closed forceps, or taking some fat or other tissue with the IFM and removing the fat as described in step 2.10.</li>
	<li>Quality control the IFM muscle sample, using the forceps to remove non-IFM muscles, fat, cuticle, etc. from the sample (Figure 2K,L). 
	NOTE: With Mef2-Gal4, IFM is labeled more strongly than other muscle types at early timepoints (Figure 2K,K&#39;), allowing removal of jump muscle and larval muscles based on fluorescence intensity and muscle shape. Fat and cuticle tissue look different and are not labeled by a muscle-specific fluorescence label (Figure 2K,K&#39;). See the discussion section for other Gal4 lines that label IFM.</li>
	<li>Using a clipped pipette tip, transfer the pile of IFMs into a 1.5 mL microcentrifuge tube filled with 250 &#956;L of chilled 1x PBS (Figure 2M-O). Proceed immediately to section 4. 
	NOTE: IFM samples may be lost simply by sticking to the side of the pipette tip. Pipetting buffer up and down several times before collecting IFMs can make standard tips less sticky, and siliconized or perfluoroalkoxy (PFA) tips (see Table of Materials) with lower surface tensions can help prevent sample loss.</li>
</ol>
3. IFM Dissection After 48 h APF
<ol>
	<li>Assemble necessary equipment including two #5 biology grade forceps, fine scissors, standard glass microscope slides, double-stick tape, pipette, pipette tips, dry ice, and (for RNA applications) isolation reagent (see Table of Materials). Chill the 1x PBS and microcentrifuge tubes on ice.</li>
	<li>Using a lightly wetted paintbrush, transfer the staged pupae to a strip of double-sided sticky tape mounted on a microscope slide (Figure 3A). Place the pupae in a line oriented in the same orientation (ventral down and anterior towards the bottom of the slide). 
	NOTE: Be careful not to use too much water on the paintbrush or filter, or the pupae will not stick well. If pupae do not stick, dry them by first transferring to a dry filter or tissue paper. Mount as many pupae as can be dissected within a 30 min time window, ideally ~10 pupae.</li>
	<li>Remove the pupa from the pupal case. Use forceps to tease apart and open the pupal case above the anterior spiracles (Figure 3B).</li>
	<li>Gently slide a pair of forceps dorsally towards the posterior, cutting the pupal case as the forceps move (Figure 3B&#39;). Be careful not to rupture the underlying pupa. Liberate the pupa from the opened case and immediately transfer it to a drop of 1x PBS on a second microscope slide (Figure 3B&#34;,C).</li>
	<li>Repeat steps 3.3 and 3.4 for all pupae in the line, then set the double-stick tape slide aside.</li>
	<li>Using the fine scissors, cut the abdomen of the pupa away from the thorax and push it into a separate pile (Figure 3D,D&#39;). Repeat for the remaining pupae. 
	NOTE: Begin timing the length of dissection with step 3.6, as soon as pupal integrity is disrupted. Dissect as many flies as possible in 20&#8211;30 min to prevent cell death and associated transcriptomic and proteomic changes. When dissecting 1 d adults or &#62;90 h pupae, it is often convenient for later steps to additionally remove the head with the fine scissors.</li>
	<li>Using a tissue paper, remove the majority of the 1x PBS (generally cloudy with suspended fat) as well as the pile of abdomens (Figure 3E). Add a drop of fresh, chilled 1x PBS to the remaining thoraxes.</li>
	<li>Use the scissors to cut the thorax in half (Figure 3F,F&#39;) by cutting from the head down the longitudinal body axis in a single motion. Alternately, if the head has been removed, first insert the scissors where the head was attached and cut the top half of the thorax longitudinally between the IFMs. Then, cut the ventral side of the thorax with a second cut in the same orientation.</li>
	<li>Repeat steps 3.7 and 3.8 for all pupae to be dissected, generating a pile of thorax hemisections near the center of the slide. Ensure there is enough chilled 1x PBS on the slide so that the hemisections do not dry out. 
	NOTE: After 48 h APF, IFMs are large enough to be visible under a standard dissecting microscope to the trained eye. At this point in the protocol, muscles with a fluorescent label can be moved to a fluorescent dissecting scope to aid in IFM identification or for training purposes, but this is not necessary.</li>
	<li>Dissect the IFMs out of the thorax. Isolate one of the hemisections using the #5 forceps (Figure 3G,H). Gently insert the tips of one forceps above and below the middle of the IFMs (Figure 3G&#39;,H&#39;). While holding the first forceps still, use fine scissors to cut one end of the IFM away from the cuticle and tendons. Then, cut the other end of the IFM free from the cuticle (Figure 3G&#39;&#39;,H&#39;&#39;). 
	NOTE: Depending on the orientation of the thorax after the first IFM cut, it is useful to rotate the thorax 180&#176; so that the second IFM cut is easier to perform.</li>
	<li>Remove the IFM bundle from the thorax with forceps (Figure 3G&#39;&#39;&#39;,H&#39;&#39;&#39;), transferring it to the edge of the PBS bubble to use water tension to hold it in place (Figure 3I). Push the carcass to the opposite side of the slide. Repeat for the remaining thorax hemisections, generating a collection of dissected IFMs. 
	NOTE: If the IFMs do not stay in a neat pile, remove some of the 1x PBS with a tissue. Be careful not to let all of the PBS evaporate, and ensure that the dissected IFMs and hemithoraxes remain covered by buffer.</li>
	<li>After dissecting all IFMs, quickly perform a quality control on the dissected muscle. Using #5 forceps, remove any jump muscle or cuticle fragments that may have found their way into the sample (Figure 3J-K&#39;&#39;). 
	NOTE: Jump muscle appears different from IFM. If dissecting Mef2-Gal4 labeled muscle under fluorescence, jump muscle has a weaker fluorescence and a different shape and texture. Under normal light, it appears nearly translucent while the IFMs are an opaque, milky yellow (Figure 3J-J&#39;&#39;,K).</li>
	<li>Using water tension, capture (but do not squish) the dissected IFMs between a pair of forceps (Figure 3L). Transfer the IFMs to a 1.5 mL microcentrifuge tube pre-filled with 250 &#956;L of chilled 1x PBS (Figure 3M). Proceed immediately with section 4. 
	NOTE: When forceps tips are brought into proximity of each other and lifted out of a buffer solution, water tension causes a bubble of buffer to be captured between the forceps tips. If IFMs are also present in this bubble, they can be lifted out of the solution and easily transferred to another buffer-filled receptacle. It is important to squeeze the forceps to bring the tips near one another without touching each other, to avoid macerating the tissue captured in the buffer bubble.</li>
</ol>
4. Pellet and Preserve the IFM Sample
<ol>
	<li>Pellet the IFMs by centrifuging the 1.5 mL microcentrifuge tube for 3&#8211;5 min at 2,000 x g in a table-top centrifuge (Figure 4A,B).</li>
	<li>Remove the buffer using a pipette tip (Figure 4C).</li>
	<li>For RNA applications, resuspend the IFM pellet in 50&#8211;100 &#956;L of the desired RNA isolation buffer (see Table of Materials, Figure 4D). Otherwise, proceed to step 4.4. 
	NOTE: IFMs can be dry-frozen after step 4.2 for mass spectrometry preparations or isolation of RNA with commercial kits (see representative results). For RNA applications, better results are obtained by immediately resuspending and freezing the IFM pellet in isolation buffer.</li>
	<li>Freeze sample on dry ice or snap freeze in liquid nitrogen (Figure 4E). Store at -80 &#176;C until ready for subsequent steps in sample preparation for downstream analysis. 
	NOTE: After cryopreservation, samples can be stored for several months before processing for downstream investigation.</li>
</ol>

The authors have nothing to disclose.

In this protocol, we present the basic technique to dissect Drosophila IFMs from early and late-stage pupae for downstream isolation of protein, DNA, RNA or other macromolecules. The protocol can be easily adapted to dissect IFM from adult flies. We demonstrate the utility of our dissection protocol for mRNA-Seq, proteomics and RT-PCR applications. With the continuous improvement of omics technologies to allow analysis of samples with less starting material and lower input concentrations, these dissections will likely become valuable for many additional applications. As IFMs are an established model for human myopathies4,24 and muscle-type specific development9,12, we envision, for example, IFM-enriched metabolomics, investigations of chromatin conformation via 3C or 4C, splicing network evaluation via CLiP interactions or phospho-proteomics of myofibrillogenesis.
It is important to consider that these dissections produce a sample enriched for IFM instead of a pure IFM sample. This is unavoidable due to motor neuron innervation, tendon attachments and tracheal invasion of muscle fibers. Bioinformatics analysis can be used to identify IFM enriched genes or proteins, but further experiments are required to demonstrate that they are in fact IFM-specific. Sample purity can be assayed using published tissue-specific markers such as Stripe45 (tendon), Act79B4,44 (tubular muscle), Act88F15 (IFM), or syb46 (neuronal specific). It may be possible to use such markers to normalize datasets to the IFM-specific content, but users are cautioned that temporal changes in expression of genes used for normalization, for example of IFM-specific genes or tubulin, may bias such an approach.
Genetically encoded tissue-specific labeling methods, for example EC-tagging47,48 or PABP-labeling49,50 for isolating RNA have been developed in recent years, which may help obtain a truly tissue-specific RNA sample. However, EC-tagging requires constant feeding of flies47 and thus is not applicable during pupal stages. The sensitivity and completeness of PABP-labeled transcriptomes may have limitations51. FACS approaches to isolate individual muscle fibers are complicated by the large size and syncytial nature of IFMs. INTACT52,53 style approaches may be applied to isolate specific subcellular-compartments from IFMs, which may prove useful for isolating pure populations of IFM nuclei or mitochondria. Manual dissections are still the current standard to obtain intact IFM tissue for most downstream applications.
Sample quality depends on several critical steps in the dissection process. The dissections are technically demanding, with dissection speed and sample purity increasing with experience. Dissecting for short periods of time (20&#8211;30 min) in chilled buffer without detergent and immediately freezing helps to preserve sample integrity, as has been observed previously for mouse tendon isolation54. IFMs can be successfully dry-frozen after removing all buffer from the pellet, but specifically for RNA isolation, freezing samples in isolation buffer tends to produce better results. IFMs from up to 20 separate dissections are combined prior to RNA or protein isolation, allowing scaling up and collecting enough material, even from early timepoints or mutants16,32, for downstream analysis.
For RNA applications, the most critical step may be the isolation of the RNA itself. Guanidinium thiocyanate-phenol-chloroform isolation (method 1 above) outperforms most commercial kits tested and, as previously noted, is considerably less expensive55. The variability observed in RNA isolation yields with commercial kits is in agreement with previous observations56,57. We further add glycogen during isopropanol precipitation to help recover all RNA. Beyond RNA yield, it is important to verify RNA integrity to ensure that the sample has not been fragmented or degraded during the dissection and isolation processes. It is also essential to work RNase-free. Lastly, the choice of RT-kit can impact the sensitivity of the reverse transcription process. While not often discussed in detail, all of these points influence the quality of the IFM sample and the data obtained from downstream applications.
Several important modifications set the protocol apart from existing IFM dissection protocols. Although a detailed dissection protocol for IFM immunofluorescence exists19, this protocol presents a different approach to pupal dissections that allows more rapid isolation of IFM tissue. This allows collection of large amounts of IFM tissue (relatively speaking) with limited dissection times to prevent proteome or transcriptome changes. Other protocols describe dissection of adult IFM for visualizing GFP staining in individual myofibrils39 or for staining of larval body-wall muscles58, but they do not address dissection at pupal stages or for isolation of RNA or protein. This approach is also distinct from the existing protocol for microdissection of pupal IFMs from cryosections38, which may generate a purer IFM sample but is more labor intensive and produces less material. As compared to other rapid adult IFM dissection protocols38,39, IFMs are isolated in PBS buffer without detergent to limit stress induction and other major expression changes.
The key advance in this protocol is the inclusion of a live, fluorescent reporter, allowing isolation of the IFMs at early pupal stages. We standardly use Mef2-GAL459 driving either UAS-CD8::GFP or UAS-GFP::Gma60. This allows differential labelling of IFM (flight muscles are more strongly labeled and differently shaped than other pupal muscles) as well as performance of GAL4-UAS-based manipulations, for instance rescue or RNAi experiments. It is also possible to combine Mef2-GAL4 with tub-GAL80ts to avoid RNAi-associated early lethality or with UAS-Dcr2 to increase RNAi efficiency40.
There are additional GAL4 drivers or GFP-lines available that vary in muscle-type specificity, temporal expression pattern, and driver strength19,61 that may be used instead of Mef2-GAL4. For example, Act88F-GAL4 is first expressed around 24 h APF, so it cannot be used for earlier timepoints; however, it strongly labels IFM and may be useful to avoid RNAi-associated early lethality. Him-GFP or Act88F-GFP label IFM, again with temporal restrictions, but they avoid GAL4 dependence of marker expression and may be useful in combination with a mutant background of interest. Lists of other possible marker lines are available19. It should also be noted that use of transgenes and the GAL4/UAS system may cause gene expression artifacts, so it is important to use appropriate controls, for example the driver line crossed to the wild-type background strain, so that such artifacts are presumably the same in all samples.
With the accompanying video, this detailed protocol aims to make pupal IFM dissection more accessible and promote the use of omics approaches to study muscle development. Coupling the power of Drosophila genetics and cell biology with the biochemistry and omics assays accessible through dissected IFM has the potential to advance mechanistic understanding of myogenesis and muscle function. Future studies linking systems-level observations of transcriptome and proteome regulation to metabolic and functional outputs will provide a deeper understanding of muscle-type specific development and the pathogenesis of muscle disorders.

Modern omics technologies provide important insights into muscle development and the mechanisms underlying human muscle disorders. For example, analysis of transcriptomics data combined with genetic and biochemical verification in animal models has revealed that loss of the splicing factor RBM20 causes dilated cardiomyopathy due to its regulation of a target network of more than 30 sarcomere genes previously associated with heart disease, including titin1,2,3.
In a second example, studies from cell culture, animal models, and human patients have shown that myotonic dystrophy is caused by a disruption in RNA regulation due to sequestration of Muscleblind (MBNL) and upregulation of CELF14,5. The cross-regulatory and temporal dynamics between MBNL and CELF1 (also called CUGBP1 or Bruno-Like 2) help&#160;to explain the persistent embryonic splicing patterns in myotonic dystrophy patients. Additionally, the large network of misregulated targets helps to explain the complex nature of the disease4,6,7,8. A majority of such studies utilize omics approaches in genetic model organisms to understand the mechanisms underlying human muscle disease. Furthermore, they highlight the importance of first understanding temporal and tissue-type specific gene expression, protein modification, and metabolic patterns in healthy muscle to understand alterations in diseased or aging muscle.
Drosophila melanogaster is another well-established genetic model organism. The structure of the sarcomere as well as individual sarcomere components are highly conserved from flies to vertebrates4,9,10, and the indirect flight muscles (IFMs) have become a powerful model to study multiple aspects of muscle development11,12. First, the fibrillar flight muscles are functionally and morphologically distinct from tubular body muscles11,13, allowing investigation of muscle-type specific developmental mechanisms. Transcription factors including Spalt major (Salm)14, Extradenticle (Exd), and Homothorax (Hth)15 have been identified as fibrillar fate regulators. Additionally, downstream of Salm, the CELF1 homolog Bruno1 (Bru1, Aret) directs a fibrillar-specific splicing program16,17.
Second, IFMs are an important model for understanding the process of myogenesis itself, from myoblast fusion and myotube attachment to myofibrillogenesis and sarcomere maturation9,18,19. Third, Drosophila genetics permits investigation of contributions by individual proteins, protein domains, and protein isoforms to sarcomere formation, function, and biophysical properties20,21,22,23. Lastly, IFM models have been developed for the study of multiple human muscle disorders, such as myotonic dystrophy, myofibrillar myopathies, muscle degenerative disorders, actinopathies, etc.24,25,26,27, and have provided important insights into disease mechanisms and potential therapies28,29,30. Thus, Drosophila is a useful model to address many open questions in the myogenesis field, including mechanisms of muscle-type specific transcription, splicing, and chromatin regulation, as well as to the role of metabolism in muscle development. The application of modern omics technologies, in particular in combination with the wide variety of genetic, biochemical and cell biological assays available in Drosophila, has the potential to dramatically advance the understanding of muscle development, aging, and disease.
IFMs are the largest muscles in the fly, spanning nearly 1 mm across the entire length of the thorax in adults31,32. However, this small size generates the challenge of obtaining enough sample to apply omics technologies in Drosophila in a tissue-type specific manner. Moreover, IFMs are part of the adult musculature that is formed during pupal stages. Myoblasts fuse to form myotubes, which attach to tendons around 24 h after puparium formation (APF) and undergo a compaction step necessary to initiate myofibrillogenesis around 30 h APF (Figure 1A-D)18,33,34.
The myofibers then grow to span the entire length of the thorax, with myofibrils undergoing an initial growth phase focused on sarcomere addition until about 48 h APF, and then transitioning to a maturation phase, in which sarcomeres grow in length and width and are remodeled to establish stretch-activation by 72 h APF (Figure 1A-D)32,35. The onset of fiber maturation is at least partially controlled by Salm and E2F32,36,37, and multiple IFM-specific sarcomere protein isoforms whose splicing is controlled by Bru1 are incorporated during this phase16,17. Mature flies eclose from 90&#8211;100 h APF. This means that to study muscle development, IFM has to be isolated with sufficient quantity, quality, and purity from multiple pupal timepoints to facilitate analysis using omics approaches.
Several protocols for IFM dissection have been published. While these protocols work well for their intended applications, none are ideal for omics approaches. Protocols that preserve IFM morphology for immunofluorescence of pupal and adult IFMs19, isolate IFM fibers for mechanical evaluation31, or utilize microdissection of pupal IFM from cryosections38 are too specialized and time and labor intensive to reasonably obtain sufficient amounts of IFM tissue for omics applications. Other protocols have been developed for rapid dissection of specifically adult IFM38,39, thus are not applicable to pupal stages, and use buffers that are not ideal or may be incompatible with, for example, RNA isolation. Thus, there is a need to develop new approaches to isolate pupal IFM for biochemistry or omics applications.
Here we present a protocol for the dissection of IFM during pupal stages that has&#160;been used successfully for mRNA-Seq analysis from 16 h APF through adult stages16,32. The protocol employs a green fluorescent protein (GFP) label to identify IFMs at all stages of pupal and adult development, allowing live dissection under a fluorescent dissecting microscope. The approach is less labor-intensive, with a higher throughput than existing IFM dissection protocols. This allows rapid isolation and cryopreservation of samples, generating enough material after several rounds of dissection for omics approaches as well as for standard reverse transcription polymerase chain reaction (RT-PCR) or western blotting.
We present the protocol in two parts, demonstrating how to rapidly dissect IFMs both before 48 h APF (during early metamorphosis, when IFM attachments are more tenuous) and after 48 h APF (when the pupal body plan and IFM attachments are well-defined). We demonstrate that we can isolate high quality RNA from dissected IFMs at all timepoints and present data on different approaches to RNA isolation and reverse transcription. Lastly, we demonstrate the application of the dissection protocol to mRNA-Seq, mass spectrometry, and RT-PCR using the CELF1 homolog Bruno1 as an example. We show misexpression of sarcomere protein isoforms in proteomics data from Bruno1 mutant IFM and examine Bruno1 regulation of the C-terminal splice event of Myosin heavy chain (Mhc). These results illustrate how omics data can provide a deeper understanding of biological phenomena, complementing genetic and biochemical experiments.

<table><tbody><tr><td>5x High Fidelity (HF) buffer</td><td>Thermo Fisher</td><td>F518L</td><td></td></tr><tr><td>60 mm culture dishes</td><td>Sigma-Aldrich</td><td>Z643084-600EA</td><td>Greiner dishes, 60 mm x 15 mM, vented</td></tr><tr><td>Black dissecting dish (glass)</td><td>Augusta Laborbedarf</td><td>42021010</td><td>Lymphbecken, black glass, 4 cm x 4 cm</td></tr><tr><td>Black silicon dissecting dishes: activated charcoal powder</td><td>Sigma-Aldrich</td><td>C9157</td><td>Also available from most pharmacies</td></tr><tr><td>Black silicon dissecting dishes: Sylgard 184</td><td>Sigma-Aldrich</td><td>761036</td><td>Dishes are made by mixing the Sylgard (~50g) with activated charcoal powder (200 mg) and curing it in Petri dishes (~4 60 mm dishes).</td></tr><tr><td>Blue pestle</td><td>Sigma-Aldrich</td><td>Z359947-100EA</td><td>Any pellet pestle that can sterilized, also can be used with a motor-driven grinder</td></tr><tr><td>Cell phone camera, Samsung Galaxy S9</td><td>Samsung</td><td>SM-G960F/DS</td><td>used for photos not taken under a microscope</td></tr><tr><td>Chloroform</td><td>PanReac AppliChem</td><td>A3691,0500</td><td></td></tr><tr><td>Confocal microscope, Leica SP8X upright confocal</td><td>Leica</td><td></td><td>www.leica-microsystems.com</td></tr><tr><td>Confocal microscope, Zeiss LSM 780 inverted confocal</td><td>Zeiss</td><td></td><td>www.zeiss.com</td></tr><tr><td>Double stick tape</td><td>Scotch/3M</td><td>3M ID 70005108587</td><td>Double-sided tape, available at most office supply handlers</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 mm x 0.02 mm tip</td></tr><tr><td>EtOH (100%, RNase free)</td><td>Sigma-Aldrich</td><td>32205-M</td><td></td></tr><tr><td>fluorescent dissecting microscope camera, Leica DFC310 FX camera</td><td>Leica</td><td></td><td>www.leica-microsystems.com</td></tr><tr><td>Fluorescent dissecting microscope software, Leica Application Suite (LAS) version 4.0.0</td><td>Leica</td><td></td><td>www.leica-microsystems.com</td></tr><tr><td>Fluorescent dissecting microscope, Leica M165 FC</td><td>Leica</td><td></td><td>www.leica-microsystems.com</td></tr><tr><td>Fly: Bru1[M2]</td><td></td><td></td><td>Fly stock; This paper</td></tr><tr><td>Fly: Bru1[M3]</td><td></td><td></td><td>Fly stock; This paper</td></tr><tr><td>Fly: Mef2-GAL4</td><td>Bloomington Stock Center</td><td>BDSC:27390</td><td>Fly stock</td></tr><tr><td>Fly: salm[1]</td><td>Bloomington Stock Center</td><td>3274</td><td>Fly stock</td></tr><tr><td>Fly: salm[FRT]</td><td></td><td></td><td>Fly stock; see Spletter et al., Elife, 2018</td></tr><tr><td>Fly: UAS-Bru1IR</td><td>Vienna Drosophila Research Center</td><td>GD41568</td><td>Fly stock, RNAi hairpin</td></tr><tr><td>Fly: UAS-GFP::Gma</td><td>Bloomington Stock Center</td><td>BDSC:31776</td><td>Fly stock</td></tr><tr><td>Fly: UAS-mCD8a::GFP</td><td>Bloomington Stock Center</td><td>BDSC:5130</td><td>Fly stock</td></tr><tr><td>Fly: w[1118]</td><td>Bloomington Stock Center</td><td>3605</td><td>Fly stock</td></tr><tr><td>Fly: weeP26</td><td></td><td></td><td>Fly stock; see Clyne et al., Genetics, 2003</td></tr><tr><td>GFP detection reagent, GFP-Booster</td><td>ChromoTek</td><td>gba488-100</td><td></td></tr><tr><td>Glycogen</td><td>Invitrogen</td><td>10814-010</td><td></td></tr><tr><td>Image processing software, Photoshop CS6</td><td>Adobe</td><td></td><td>www.adobe.com</td></tr><tr><td>Isopropanol</td><td>Sigma-Aldrich</td><td>I9516-25ML</td><td>2-propanol</td></tr><tr><td>Method 1 (RNA isolation): TRIzol</td><td>Life Technologies</td><td>15596018</td><td>Guanidinium isothiocyanate and phenol monophasic solution</td></tr><tr><td>Method 2 (RNA isolation): Method 1 + TURBO DNA-free Kit</td><td>Invitrogen</td><td>AM1907</td><td>TRIzol isolation followed by treatment with a kit to remove DNA</td></tr><tr><td>Method 3 (RNA isolation): Direct-zol RNA Miniprep Plus Kit</td><td>Zymo Research</td><td>R2070S</td><td>RNA isolation in TRIzol, but over commercial columns instead of using phase separation. Recommended DNase treatment performed with Monarch Dnase I in Monarch DNase I Reaction buffer.</td></tr><tr><td>Method 4 (RNA isolation): RNeasy Plus Mini Kit</td><td>Qiagen</td><td>74134</td><td>We used the provided DNase treatment. IFM pellets were homogenized in RTL buffer as suggested for animal tissues.</td></tr><tr><td>Method 5 (RNA isolation): ReliaPrep RNA Tissue Miniprep System</td><td>Promega</td><td>Z6110</td><td>We applied the protocol for &amp;lsquo;Purification of RNA from Fibrous Tissues&amp;rsquo;.</td></tr><tr><td>Method 6 (RNA isolation): Monarch Total RNA Miniprep Kit</td><td>New England Biolabs</td><td>T2010G</td><td>We applied the protocol for tissues/leukocytes and lysed in 300 &amp;mu;L of RNA Protection Reagent.</td></tr><tr><td>Microcentrifuge tubes</td><td>Thermo Fisher</td><td>AM12400</td><td>RNase-free Microfuge Tubes, 1.5 mL</td></tr><tr><td>Microscope slides</td><td>Thermo Fisher</td><td>12342108</td><td>Standard slides, uncharged, 1 mm</td></tr><tr><td>Microtome blades</td><td>PFM Medical</td><td>207500003</td><td>C35 feather 80 mm</td></tr><tr><td>Monarch DNase I</td><td>New England Biolabs</td><td>T2004-21</td><td></td></tr><tr><td>Monarch DNase I Reaction Buffer</td><td>New England Biolabs</td><td>T2005-21</td><td></td></tr><tr><td>Normal goat serum</td><td>Thermo Fisher</td><td>16210072</td><td></td></tr><tr><td>OneTaq Polymerase</td><td>New England Biolabs</td><td>M0480X</td><td></td></tr><tr><td>Paintbrush</td><td>Marabu</td><td>1910000000</td><td>Marabu Fino Round No. 0, or similar brush from any art supply</td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich</td><td>158127</td><td></td></tr><tr><td>PBS buffer (1x)</td><td>Sigma-Aldrich</td><td>P4417</td><td>Phosphate buffered saline tablets for 1 L solutions, pH 7.4</td></tr><tr><td>PFA PureTip Pipette Tips</td><td>Elemental Scientific</td><td>ES-7000-0101</td><td>Optional substitute for standard pipette tips to reduce sample loss; 100 mL, 0.8 mm orifice</td></tr><tr><td>Phusion High Fidelity Polymerase</td><td>Thermo Fisher</td><td>F-530XL</td><td></td></tr><tr><td>Pipette tips</td><td>Sigma-Aldrich</td><td>P5161</td><td>Universal 200 mL pipette tips</td></tr><tr><td>Preomics iST 8x Kit</td><td>Preomics</td><td>P.O.00001</td><td>peptide preparation kit for mass spectrometry</td></tr><tr><td>Q Exactive mass spectrometer</td><td>Thermo Fisher</td><td>725500</td><td>mass spectrometry was performed at the Protein Analysis Unit of the LMU Biomedical Center</td></tr><tr><td>Qubit RNA Assay Kit</td><td>Life Technologies</td><td>Q32855</td><td></td></tr><tr><td>Rhodamine-phalloidin</td><td>Invitrogen, Molecular Probes</td><td>10063052</td><td></td></tr><tr><td>RNA concentration Approach 1 &amp;amp; RNA integrity traces, Bioanalyzer</td><td>Agilent Technologies</td><td>G2939BA</td><td></td></tr><tr><td>RNA concentration Approach 2, Nanodrop</td><td>Thermo Fisher</td><td>ND-2000</td><td></td></tr><tr><td>RNA concentration Approach 3, Qubit 4 Fluorometer</td><td>Invitrogen</td><td>Q33238</td><td></td></tr><tr><td>RNA Pico Chips</td><td>Agilent Technologies</td><td>5067-1513</td><td></td></tr><tr><td>RNase A</td><td>Promega</td><td>A7937</td><td></td></tr><tr><td>RNase-free water, Diethyl pyrocarbonate (DEPC)</td><td>Sigma-Aldrich</td><td>D5758</td><td>DEPC treat water overnight and then autoclave, to remove all RNase.</td></tr><tr><td>RT Kit #1: Super Script III Reverse Transcriptase Kit</td><td>Invitrogen</td><td>18080-044</td><td>reverse transcription kit</td></tr><tr><td>RT Kit #2: LunaScript</td><td>New England Biolabs</td><td>E3010S</td><td>reverse transcription kit</td></tr><tr><td>RT Kit #3: QuantiNova Reverse Transcription Kit</td><td>Qiagen</td><td>205410</td><td>reverse transcription kit</td></tr><tr><td>Slide mounting buffer, Vectashield</td><td>Vector Laboratories</td><td>H-1200</td><td>containing DAPI</td></tr><tr><td>Statistical software: GraphPad Prism</td><td>GraphPad Prism</td><td></td><td>www.graphpad.com</td></tr><tr><td>Statistical software: Microscoft Excel</td><td>Microsoft</td><td></td><td>Purchased as part of the bundle: Office Home &amp;amp; Student 2019</td></tr><tr><td>Table-top centrifuge</td><td>Eppendorf</td><td>5405000760</td><td>Eppendorf Centrifuge 5425 or equivalent</td></tr><tr><td>Tissue/ Kimwipes</td><td>Sigma-Aldrich</td><td>Z188956</td><td>Standard tissue wipes</td></tr><tr><td>Transfer pipette</td><td>Sigma-Aldrich</td><td>Z350796</td><td>Plastic pipette</td></tr><tr><td>Triton-X100</td><td>Sigma-Aldrich</td><td>T9284-500ML</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>Whatman paper</td><td>Sigma-Aldrich</td><td>1004-070</td><td>Filter paper circles, Grade 4, 70 mm</td></tr></tbody></table>

dissection of drosophila melanogaster flight muscles for omics approaches

Drosophila flight muscle is a powerful model to study diverse processes such as transcriptional regulation, alternative splicing, metabolism, and mechanobiology, which all influence muscle development and myofibrillogenesis. Omics data, such as those generated by mass spectrometry or deep sequencing, can provide important mechanistic insights into these biological processes. For such approaches, it is beneficial to analyze tissue-specific samples to increase both selectivity and specificity of the omics fingerprints. Here we present a protocol for dissection of fluorescent-labeled flight muscle from live pupae to generate highly enriched muscle samples for omics applications. We first describe how to dissect flight muscles at early pupal stages (&#60;48 h after puparium formation [APF]), when the muscles are discernable by green fluorescent protein (GFP) labeling. We then describe how to dissect muscles from late pupae (&#62;48 h APF) or adults, when muscles are distinguishable under a dissecting microscope. The accompanying video protocol will make these technically demanding dissections more widely accessible to the muscle and Drosophila research communities. For RNA applications, we assay the quantity and quality of RNA that can be isolated at different time points and with different approaches. We further show that Bruno1 (Bru1) is necessary for a temporal shift in myosin heavy chain (Mhc) splicing, demonstrating that dissected muscles can be used for mRNA-Seq, mass spectrometry, and reverse transcription polymerase chain reaction (RT-PCR) applications. This dissection protocol will help promote tissue-specific omics analyses and can be generally applied to study multiple biological aspects of myogenesis.

Watch this Scientific Journal Video about Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches at JoVE.com

Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

Drosophila flight muscle is a powerful model to study transcriptional regulation, alternative splicing, metabolism, and mechanobiology. We present a protocol for dissection of fluorescent-labeled flight muscle from live pupae to generate highly enriched samples ideal for proteomics and deep-sequencing. These samples can offer important mechanistic insights into diverse aspects of muscle development.

Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

Drosophila flight muscle is a powerful model to study diverse processes such as transcriptional regulation, alternative splicing, metabolism, and ...

dissection-drosophila-melanogaster-flight-muscles-for-omics

Nanyang Technological University

Research

JoVE Journal

Genetics

11.8K Views.  Ludwig-Maximilians-University Munich. Het protocol vergemakkelijkt de dissectie van GFP-gelabelde indirecte vluchtspier van Drosophila op verschillende tijdspunten tijdens de ontwikkeling van poppen om veranderingen in gen- en eiwitexpressie te beoordelen. Met behulp van deze technieken kan een hoog verrijkt vluchtspiermonster met minimale RNA-eiwitdegradatie worden bereikt dat geschikt is voor RT-PCR, westelijke vlek of omische benaderingen. Deze gespecialiseerde ontledingen vereisen oefening en vaardigheid, maar het aantal popp...

Video: Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Neuroscience

Drosophila Larval NMJ Dissection

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 35 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. In order to access the NMJ for study, it is necessary to carefully dissect each larva. In this article we demonstrate how to properly dissect Drosophila larvae for study of the NMJ by removing all internal organs while leaving the body wall intact. This technique is suitable to prepare larvae for imaging, immunohistochemistry, or electrophysiology.

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

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Biology

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

Detection and Quantification of Single mRNA Molecules in Drosophila

Studying Muscle Transcriptional Dynamics at Single-molecule Scales in Drosophila

Skeletal muscles are large syncytia made up of many bundled myofibers that produce forces and enable body motion. Drosophila is a classical model to study muscle biology. The combination of both Drosophila genetics and advanced omics approaches led to the identification of key conserved molecules that regulate muscle morphogenesis and regeneration. However, the transcriptional dynamics of these molecules and the spatial distribution of their messenger RNA within the syncytia cannot be assessed by conventional methods. Here we optimized an existing single-molecule RNA fluorescence in situ hybridization (smFISH) method to enable the detection and quantification of individual mRNA molecules within adult flight muscles and their muscle stem cells. As a proof of concept, we have analyzed the mRNA expression and distribution of two evolutionary conserved transcription factors, Mef2 and Zfh1/Zeb. We show that this method can efficiently detect and quantify single mRNA molecules for both transcripts in the muscle precursor cells, adult muscles, and muscle stem cells.

Author Spotlight: Investigating mRNA Spatial Distribution in Drosophila Muscle Tissue 

Drosophila is a well-established model for studying key molecules that regulate myogenesis. However, current methods are insufficient to determine mRNA transcriptional dynamics and spatial distribution within syncytia. To address this limitation, we optimized an RNA fluorescence in situ hybridization method allowing the detection and quantification of mRNAs at a single-molecule scale.

Skeletal muscles are large syncytia made up of many bundled myofibers that produce forces and enable body motion. Drosophila is a classical model to study ...

Developmental Biology

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in communication with the epidermal cells they innervate and thus their analysis benefits from in situ visualization of both neuronally and epidermally expressed proteins by immunofluorescence. Traditional methods of preparing larval fillets for immunofluorescence experiments leave intact the muscle tissue that covers most of the body wall, presenting several challenges to imaging neuronal and epidermal proteins. Here we describe a method for removing muscle tissue from Drosophila larval fillets. This protocol enables imaging of proteins that are otherwise obscured by muscle tissue, improves signal to noise ratio, and facilitates the use of super-resolution microscopy to study da neuron development.

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Studies of neuronal morphogenesis using Drosophila larval dendritic arborization (da) neurons benefit from in situ visualization of neuronal and epidermal proteins by immunofluorescence. We describe a procedure that improves immunofluorescence analysis of da neurons and surrounding epidermal cells by removing muscle tissue from the larval body wall.

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in ...

Dissection and Immunohistochemistry of the Drosophila Adult Leg to Detect Changes at the Neuromuscular Junction for an Identified Motor Neuron

Drosophila melanogaster represents a genetically tractable model to study neuronal structure and function, and subsequent changes in disease states. The well characterized larval neuromuscular junction is often used for such studies. However, rapid larval development followed by muscle histolysis and nervous system remodeling during metamorphosis makes this model problematic for the study of slow age-dependent degenerative changes like those occurring in amyotrophic lateral sclerosis. Alternatively, adult flies live for 90 days and the adult leg can be used to study motor neuron changes over the course of adult lifespan using in vivo fluorescent imaging through the cuticle. Here, we describe a leg dissection technique coupled with immunocytochemistry, which allows for the study of molecular changes at the neuromuscular junction of identified adult leg motor neurons. These techniques can be coupled with a myriad of antibodies labeling both pre- and post-synaptic structures. Together these procedures allow a more complete characterization of slow age-dependent changes in adult flies and can be applied across multiple motor neuron disease models.

Dissection and Immunohistochemistry of the Drosophila Adult Leg to Detect Changes at the Neuromuscular Junction for an Identified Motor Neuron

We describe a dissection technique that preserves the architecture of the neuromuscular junction and enables a detailed immunocytochemical study of motor neurons in the adult Drosophila leg.

Drosophila melanogaster represents a genetically tractable model to study neuronal structure and function, and subsequent changes in disease states. The ...

Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles

Muscle mass wasting, known as muscle atrophy, is a common phenotype in Drosophila models of neuromuscular diseases. We have used the indirect flight muscles (IFMs) of flies, specifically the dorso-longitudinal muscles (DLM), as the experimental subject to measure the atrophic phenotype brought about by different genetic causes. In this protocol, we describe how to embed fly thorax muscles for semi thin sectioning, how to obtain a good contrast between muscle and the surrounding tissue, and how to process optical microscope images for semiautomatic acquisition of quantifiable data and analysis. We describe three specific applications of the methodological pipeline. First, we show how the method can be applied to quantify muscle degeneration in a myotonic dystrophy fly model; second, measurement of muscle cross-sectional area can help to identify genes that either promote or prevent muscle atrophy and/or muscle degeneration; third, this protocol can be applied to determine whether a candidate compound is able to significantly modify a given atrophic phenotype induced by a disease-causing mutation or by an environmental trigger.

Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles

We report a method to quantify muscle area, which is an indirect method to determine muscle mass in Drosophila adults. We demonstrate the application of our methodology by analyzing the indirect flight muscles in a Drosophila model of Myotonic Dystrophy disease.

Muscle mass wasting, known as muscle atrophy, is a common phenotype in Drosophila models of neuromuscular diseases. We have used the indirect flight ...

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique cristae-rich double membrane ultrastructure of this organelle is elegantly arranged to carry out multiple functions by partitioning biomolecules. Mitochondrial ultrastructure is intimately linked with various functions; however, the fine details of these structure-function relationships are only beginning to be described. Here, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle. Serial-section electron tomography may be adapted to study any cellular structure in three-dimensions.

3D Mitochondrial Ultrastructure of Drosophila Indirect Flight Muscle Revealed by Serial-section Electron Tomography

In this protocol, we demonstrate the application of serial-section electron tomography to elucidate mitochondrial structure in Drosophila indirect flight muscle.

Mitochondria are cellular powerhouses that produce ATP, lipids, and metabolites, as well as regulate calcium homeostasis and cell death. The unique ...

Assessing the Structural Integrity of the Dorsal Longitudinal Muscle Neuromuscular Junctions in Drosophila

Source: Sidisky, J. M., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/61363">Visualizing Synaptic Degeneration in Adult Drosophila in Association with Neurodegeneration.</a> J. Vis. Exp. (2020).
This video demonstrates a protocol for assessing the structural integrity of neuromuscular junctions in the dorsal longitudinal muscles of Drosophila. The procedure involves staining neurons and muscles with two distinct antibodies and comparing fluorescence patterns between mutant and wild-type flies. A reduction in neuronal staining in mutant flies indicates neurodegeneration, while the muscle structure remains intact.

1. Flash freezing and thorax bisection

	Before beginning the bisections, fill a Dewar flask with liquid nitrogen wearing proper cryo-protective gloves ...

JoVE Encyclopedia of Experiments

Neuropathology

Neuromuscular Disorders

Drosophila Late Pupa Indirect Flight Muscle (IFM) Dissection: A Method for High-Throughput Tissue Collection

Source: Kao, S. Y., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/60309/" target="_blank">Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches</a>. J. Vis. Exp. (2019).
The Drosophila flight muscles are used as a model system for muscle development and physiology. This video describes the flight musculature of adults and highlights a protocol to dissect developing indirect flight muscles that are suitable for RNA sequencing from pupae.

Source: Kao, S. Y., et al. Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches. J. Vis. Exp. (2019).
The Drosophila flight muscles ...

Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

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