We thank Alain Vincent for his critical reading of the manuscript. We thank Ruth Lehmann for providing the Zfh1 antibody. We acknowledge the help of Stephanie Dutertre and Xavier Pinson for confocal microscopy at the MRic Imaging Platform. EL is supported by PhD fellowship from minist&#232;re de l&#39;enseignement sup&#233;rieur et de la recherche scientifique. NA is supported by the AFM-Telethon Ph.D. fellowship 23846. TP is funded by a Chan Zuckerberg Initiative DAF grant to T.P. (2019-198009). HB is supported by CNRS, the Atip Avenir program, and the AFM-Telethon trampoline grant 23108.

1. Wing disc and adult muscle dissection and preparation
NOTE: Clean up the dissection bench and all the dissection instruments with RNAse inhibitor solution (Table of Materials). Laboratory gloves should be worn during the whole experimental procedure.
<ol>
	<li>Wing disc
	<ol>
		<li>Place the L3 staged larvae in cold 1x phosphate-buffered saline (PBS) (Figure 1A).</li>
		<li>Rinse the larvae 3x with 1x PBS and keep them in this solution for 5 min.</li>
		<li>Using two pairs of sharp forceps, hold the larva from the anterior part and pull and discard the rest of the body tissues (around 2/3).</li>
		<li>Hold both the first 1/3 of the larva with one pair of forceps and the mouth hooks with the other pair. Retract the mouth hooks inside the larva until it is fully inverted. 
		NOTE: Dissect approximately 30 larvae per experiment, to ensure adequate sample size for analysis.</li>
		<li>Observe that the wing discs are closely associated with the two primary branches of the tracheae, which run along each side of the larva.</li>
		<li>Remove all the other larval components (leg discs and brain) to obtain a clean inverted larval carcass with a pair of wing discs attached to the trachea (Figure 1B).</li>
		<li>Place the samples in a 1 mL centrifuge tube and fix them in 4% paraformaldehyde (PFA) diluted in 1x PBS for 45 min at room temperature under agitation. 
		NOTE: The sample agitation is achieved by an oscillating stirrer (Table of Materials).</li>
		<li>Wash the samples 3 x 5 min with 70% ethanol (EtOH, diluted in RNAse-free water) and store them in the same washing solution at 4 &#176;C for at least 2 days and up to 1 week.</li>
	</ol>
	</li>
	<li>Indirect flight muscles
	<ol>
		<li>Anesthetize the adult flies on a CO2 fly pad and remove the heads, abdomens, legs, and wings.</li>
		<li>Place the dissected thoraxes in cold 1x PBS and continue doing so until all samples are dissected (Figure 1D).</li>
		<li>Prefix the thoraxes in 4% PFA, (1% Triton) for 20 min at room temperature under agitation.</li>
		<li>Rinse the samples 3 x 5 min with 1x PBT washing solution (1% Triton in 1x PBS).</li>
		<li>Position the thoraxes on a double-sided tape (Table of Materials) on a glass slide and bisect them with a sharp microtome blade to produce two hemi-thoraces (Figure 1E,F).</li>
		<li>Fix the samples in 4% PFA (1% Triton) for 45 min at room temperature under agitation.</li>
		<li>Wash 2 x 20 min with 1x PBT washing solution (1% Triton, 1x PBS) at room temperature under agitation.</li>
		<li>Place the hemi-thoraces in cold 1x PBS and use the forceps to gently isolate the IFMs from the hemi-thorace (Figure 1G). 
		&#8203;NOTE: To achieve this, it is important to remove the cuticle as thoroughly as possible.</li>
		<li>To permeabilize the samples, store the isolated IFMs in 70% EtOH solution for at least 2 days and up to 1 week at 4 &#176;C. 
		&#8203;NOTE: Isolate the IFMs from approximately 10 thoraxes per experiment to ensure adequate sample size for analysis.</li>
	</ol>
	</li>
</ol>
2. Hybridization, immunostaining, and mounting
NOTE: Similar hybridization procedure is applied for both wing discs and IFMs samples. It is critical to isolate the wing discs from the larval carcasses and put them in 200 &#181;L of Buffer A prior to starting the hybridization.
<ol>
	<li>Resuspend the probes in 400 &#181;L of TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) for a final stock solution of 12.5 &#181;M.</li>
	<li>Transfer the samples to a 0.5 mL centrifuge tube containing Buffer A (Table of Materials). 
	NOTE: By decreasing the volume of the container, it becomes easier to visualize the samples and avoid losing them during the washing process.</li>
	<li>Wash the samples 2 x 20 min with Buffer A.</li>
	<li>Aspirate Buffer A, replace it with 400 &#181;L of the hybridization buffer (Table of Materials), and prehybridize the samples for 30 min at 37 &#176;C in a thermal mixer (Table of Materials).</li>
	<li>Dilute the probes to a final concentration of 125 nM for wing discs and 200 nM for adult muscles, in the hybridization buffer and add the primary antibody. 
	NOTE: The antibody dilution varies depending on the specific antibody being used (Table of Materials). Adding antibodies is optional. Users may opt to forego protein staining and instead use a similar smFISH protocol that does not include the addition of antibodies.</li>
	<li>Aspirate the hybridization buffer and replace it with 100 &#181;L of hybridization buffer containing the probe and the antibody.</li>
	<li>Incubate the samples in the dark at 37 &#176;C for at least 16 h under 300 rpm agitation in a thermal mixer.</li>
	<li>Warm up an aliquot of Buffer A at 37 &#176;C.</li>
	<li>Wash the samples for 3 x 10 min in the thermal mixer at 37 &#176;C under 300 rpm agitation.</li>
	<li>During the last wash, dilute the secondary antibody (1/200) and DAPI (1/10,000) in Buffer A and incubate the samples in this solution at 37 &#176;C for at least 1 h.</li>
	<li>Wash the samples for 3 x 20 min with Buffer B (Table of Materials).</li>
	<li>Carefully transfer the samples on a microscope slide, wipe off the residual Buffer B, and add 30 &#181;L of mounting medium, and cover with an 18 x 18 mm coverslip. Use a nail polish to seal the preparation.</li>
	<li>Store the samples for up to 1 week at 4 &#176;C. 
	&#8203;NOTE: However, it is recommended to perform the imaging as soon as possible to avoid probe signal degradation.</li>
</ol>
3. Imaging
<ol>
	<li>Acquire images with an xyz acquisition mode using a confocal microscope equipped with 40x and 63x oil immersion objectives (Table of Materials). The smFISH signal is detected by the HyD detector via a photon counting mode.</li>
	<li>Locate the samples using the DAPI signal and UV lamp.</li>
	<li>To image the wing disc-associated muscle progenitors and IFMs (Figure 2 and Figure 3), apply the following setup for laser lines and emission filters: select DAPI (excitation (ex.) 405 nm, emission (em.) 450 nm), Alexa Fluor 488 (ex. 496 nm, em. 519 nm), and Quasar 670 for the smFISH probes (ex. 647 nm, em. 670 nm).</li>
	<li>To image the adult MuSCs (Figure 2D), apply the following setup for laser lines and emission filters: select DAPI (ex. 405 nm, em. 450 nm), Alexa Fluor 488 (ex. 496 nm, em. 519 nm), Alexa Fluor 555 (ex. 555 nm, em. 565 nm), and Quasar 670 for the smFISH probes (ex. 647 nm, em. 670 nm). 
	NOTE: For the imaging, the wavelengths can be adapted according to the dyes associated with secondary antibodies and probes.</li>
	<li>Save the confocal images as .TIFF files.</li>
</ol>
4. Post imaging analysis
<ol>
	<li>To get started, launch ImageJ and navigate to the Plugins menu. From there, select Macros | Edit to open the macros source code. 
	NOTE: This will enable adjustment of the parameters as needed, and if required, modify the folder directory for each channel within the macro&#39;s source code.</li>
	<li>To follow this protocol and quantify Mef2 smFISH spots in larvae, use the following settings: Log_radius = 3.0 and Log_quality = 20.0. Segement Mef2-positive nuclei with blur = 2; nucleus_scale_parameter = 30; nucleus_threshold = -8.0; nucleus_size = 300.</li>
	<li>In muscle fibers, use the following parameters: Log_radius = 2.5; Log_quality = 60.0; blur = 2; nucleus_scale_parameter = 100; nucleus_threshold = 0.0; nucleus_size = 300.</li>
	<li>To initiate the macro, simply execute the run command and wait for the macro to automatically load all images from the designated folder and quantify them in a sequential manner. 
	NOTE: Please note that the duration of this process may vary from a few seconds to several hours, depending on the volume of data being analyzed.</li>
	<li>Look for the results displayed in two new .csv files: file_FISH_results and file_nuclei_results. The first shows the number of transcriptional spots and their average and maximum intensities.&#160;The second one gives both the total number of nuclei and the number of spots inside each nucleus.</li>
</ol>

Detection and Quantification of Single mRNA Molecules in Drosophila

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The authors have no conflicts of interest to disclose.

The application of the smFISH method has gained popularity in recent times, with its use extending to various cell types and model organisms. However, in the case of Drosophila, the majority of knowledge about adult muscle gene expression is derived from bulk molecular assays, which fail to provide quantitative information on the precise spatial location of mRNA molecules. To address this gap, we have optimized a method for performing smFISH on muscle precursor cells and adult Drosophila muscles. This approach has been adapted from a previously published protocol23 and optimized for Drosophila muscle tissues.
The major obstacle in obtaining high-quality smFISH images is the thickness of adult muscles, which hinders the optimal penetration of probes. Therefore, it is crucial to separate the adult muscles from the other tissues of the animal and carry out the hybridization process with mild agitation. This particular step ensures effective permeabilization of the tissues.
Another critical aspect is that the signal-to-noise ratio can cause the computational pipeline to fail in detecting certain mRNA spots. We found that increasing the signal-to-noise ratio can be achieved by finding the optimal concentration of probes. The optimal dilution may differ depending on the composition of each probe set, including the conjugated dye and oligonucleotide composition. We recommend trying different dilutions; a final dilution of 200 nM yielded the best signal-to-noise ratio for adult tissues in these experiments.
With the smFISH method, it is possible to quantify the number of newly synthesized RNAs at the TS foci. When a gene is actively transcribed, multiple nascent RNAs are produced concurrently at the TS. As a result, the TS&#39;s intensity will surpass that of mature cytoplasmic RNA, and this feature, combined with its nuclear localization, can differentiate the TS from individual cytoplasmic RNAs. In the context of muscle biology, TS detection and quantification are particularly important for determining the synchronization of transcription of a specific gene among muscle nuclei within the same syncytium. However, this computational pipeline is not designed to differentiate between cytoplasmic mRNA and TS signals. As an alternative, we suggest combining this smFISH method with other well-established smFISH analysis tools, such as BayFISH or FISH-quant29,30. These tools have been proven to facilitate automated segmentation and fluorescence intensity calculations of RNA aggregates with remarkable precision.
Finally, while smFISH detects mRNA molecules with high spatial resolution, it is limited to analyzing a small number of mRNA at the same time. Multiscale methods like merFISH (multiplexed error-robust fluorescence in situ hybridization) enable the simultaneous analysis of a large number of different mRNA31. By combining oligonucleotide probes and error-correcting codes, this method facilitates the detection of hundreds of RNA species in single cells.

Adult skeletal muscles are made of differentiated multinucleated myofibers whose contractile properties generate movements. Muscle growth, maintenance, and regeneration rely upon muscle progenitors and muscle stem cells (MuSCs) that are specified during embryonic development1. The myogenic program is finely controlled by a set of core myogenic transcription factors (TFs) (e.g., Pax3/7, MYOD, Mef2, and ZEB)2,3,4. Deciphering the molecular mechanisms regulating muscle biology is important for elucidating fundamental myology-related questions and for possible therapeutic use in treating muscle-degenerative diseases.
Drosophila has a long history as a genetic model to study myogenesis5. It has recently emerged as a new model to investigate muscle regeneration6,7,8. The muscle structure and core myogenic programs are highly conserved between flies and mammals. For example, the TFs Mef2 and Zfh1/ZEB have a conserved function in regulating muscle development and regeneration3,9,10,11. Adult Drosophila muscles, such as the Indirect Flight Muscles (IFMs), are formed from a specific population of MuSCs, known as Adult Muscle Progenitors (AMPs)12. These AMPs are specified during embryogenesis and associate with epithelial structures such as wing and leg discs. Throughout embryonic and larval stages, AMPs remain undifferentiated until metamorphosis when they engage in differentiation and fusion to form the IFMs13,14. The TF Spalt major (spalt, salm) is expressed during flight muscle development and is necessary to determine their structural identity15. Mef2 is another important myogenic TF that is essential for adult muscle formation16,17. It is expressed in the AMPs and maintained in the adult IFMs10,18. While the majority of AMPs differentiate into functional muscles, a subset escapes differentiation and forms the adult MuSCs11. Similar to vertebrates, the TF Zfh1/ZEB is required to prevent premature differentiation of the adult MuSCs and to maintain their stemness9,10.
Gene expression dynamics have been proven to regulate various muscle biological processes19,20. The advent of high-throughput single-cell and single-nuclei RNA sequencing techniques has enabled a comprehensive exploration of these transcriptional dynamics21,22. One notable limitation of these approaches is their inability to provide the spatial distribution of mRNA molecules in the multinucleated muscle fibers. These features can be investigated by single-molecule fluorescence in situ hybridization (smFISH) that reveals two types of mRNA bodies: 1. Individual mRNA molecules spread out in the cytoplasm and representing the mature RNA and 2. A maximum of two bright nuclear foci correspond to the nascent transcript and reveal the transcriptionally active alleles23. Therefore, smFISH is a method of choice for individual mRNA molecule quantification, investigating their spatial distribution and providing snapshots of the gene transcriptional dynamics.
The smFISH method relies on a set of short fluorescent oligonucleotide probes specifically designed to be complementary to the target transcript. Upon annealing, it generates high-intensity point sources permitting the detection of mRNA at a single-molecule level using confocal microscopy24. This method is increasingly applied for a wide variety of cell types, including mammalian muscle tissues19,20. However, in Drosophila like other animal models, most of the knowledge about adult muscle gene expression is derived from bulk molecular assays, which lack quantitative information on the spatial location of mRNA molecules. Here we optimized a method for performing smFISH on muscle precursor cells and adult Drosophila muscles23,25. This protocol includes a fully automated analysis pipeline for nuclei segmentation and mRNA counting and localization.

<table><tbody><tr><td>Confocal microscope SP8</td><td>Leica</td><td>SP8 DMI 6000</td><td></td></tr><tr><td>DAPI</td><td>ThermoFisher</td><td>62248</td><td></td></tr><tr><td>Double-sided tape&amp;nbsp;</td><td>Tesa&amp;nbsp;</td><td>57910-00000</td><td></td></tr><tr><td>Dumont #55 Forceps</td><td>Fine Science Tools</td><td>11255-20</td><td></td></tr><tr><td>Formaldehyde 16%</td><td>Euromedex</td><td>EM-15710</td><td></td></tr><tr><td>Mef2 antibody</td><td>DHSB</td><td>NA</td><td></td></tr><tr><td>PBS 10x</td><td>Euromedex</td><td>ET330</td><td></td></tr><tr><td>RNaseZAP</td><td>Sigma-Aldrich</td><td>R2020</td><td></td></tr><tr><td>Stellaris probes</td><td>LGC</td><td>NA</td><td></td></tr><tr><td>Stellaris RNA FISH Hybridization Buffer</td><td>LGC</td><td>SMF-HB1-10</td><td></td></tr><tr><td>Stellaris RNA FISH Wash Buffer A</td><td>LGC</td><td>SMF-WA1-60</td><td></td></tr><tr><td>Stellaris RNA FISH Wash Buffer B</td><td>LGC</td><td>SMF-WB1-20</td><td></td></tr><tr><td>Swann-Morton Blades</td><td>Fisher scientific</td><td>0210</td><td></td></tr><tr><td>ThermoMixer</td><td>Eppendorf</td><td>5382000015</td><td></td></tr><tr><td>Triton X-100</td><td>Euromedex</td><td>2000</td><td></td></tr><tr><td>UAS-mCD8::GFP line</td><td>Bloomington</td><td>RRID:BDSC_5137</td><td></td></tr><tr><td>Ultrapure water</td><td>Sigma-Aldrich</td><td>95284</td><td></td></tr><tr><td>Watch Glass, Square, 1 5/8 in&amp;nbsp;</td><td>Carolina</td><td>742300</td><td></td></tr><tr><td>Zfh1 antibody</td><td>Gifted by Ruth Leahman</td><td></td><td></td></tr><tr><td>Zfh1-Gal4</td><td>Bloomington</td><td>BDSC: 49924, FLYB: FBtp0059625</td><td></td></tr></tbody></table>

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.

In this protocol, we used commercially produced probes targeting Mef2 and zfh1 mRNA. We designed the probes with the manufacturer&#39;s FISH probes designer (Table of Materials). The Mef2 set targets the exons common to all Mef2 RNA isoforms (from exon 3 to exon 10). The zfh1 set targets the third exon common to both zfh1-RB and zfh1-RE isoforms10. Both probes are conjugated to the Quasar 670 fluorescent dye.
We generated a macro in-house that was compatible with ImageJ software to automatically analyze the raw .tif data. The macro allows the 3D segmentation of the nuclei by Cellpose26 with the Fiji plugin BIOP and the transcriptional spot quantification by a Laplacian of Gaussian as implemented in the Trackmate plugin27. MorpholibJ plugin28 is used for postprocessing (Supplemental File 1).
As a first step, we performed smFISH on wing imaginal discs, where Mef2 and zfh1 are known to be expressed in the AMPs. These data show that both transcripts are uniformly detected in the AMPs population co-stained with either Mef2 or Zfh1 antibodies (Figure 2A,B). Active transcription sites (TS) can be revealed and distinguished from mature mRNA by smFISH as they tend to be larger and have more intense signals than individual cytoplasmic transcripts or mature nuclear transcripts. Consistently, higher magnifications of AMPs distinguish between TS foci and mature mRNA scattered in the cytoplasm (Figure 2A&#39;,B&#39;), validating the sensitivity of this smFISH protocol. It should, however, be mentioned that we observed one active TS per nucleus for both zfh1 and Mef2 (Figure 2A&#39;,B&#39;). These data further validate the accuracy of the probe design since Mef2 and zfh1 transcriptional patterns in the AMPs colocalize with the detection of their respective proteins (Figure 2).
In a second step, we examined the transcription site and distribution of Mef2 and zfh1 mRNAs in differentiated adult IFMs and associated stem cells (Figure 2C,D). These data clearly illustrate Mef2 TSs in the syncytial muscle nuclei and Mef2 mRNA distribution throughout the cytoplasm (Figure 2C,C&#39;). Zfh1 specifically marks the MuSCs population10,11. Using this smFISH protocol, we successfully detected zfh1 transcription in MuSCs marked by Zfh1-Gal4 &#62; GFP expression. Higher magnification illustrates the detection of both zfh1 TS and cytoplasmic single mRNAs (Figure 2D&#39;).
Finally, we used our in-house-built ImageJ macro to quantitatively analyze Mef2 transcriptional dynamics and spatial distribution. The computational pipeline can efficiently detect and segment Mef2 spots and muscle nuclei (Figure 3A-C). However, due to signal-to-noise ratio issues, some spots may not be detected in certain cases (Figure 3A&#39;,B&#39;). We used this automated method to investigate whether Mef2 mRNA abundance varies between muscle precursor cells (AMPs) and differentiated adult IFMs. Our analysis shows that the number of Mef2 mRNA per nuclei in adult IFMs and the AMPs are not significantly different (Figure 3D).
To gain insight into the spatial distribution of Mef2 transcripts in adult IFM muscles, we quantified the fraction of cytoplasmic versus nuclear Mef2 mRNA spots (Figure 3E,F). Using our program, we counted both the total number of Mef2 mRNA spots and the number of Mef2 mRNA spots overlapping with nuclei staining (Mef2). Subtracting the nuclei-associated mRNA from the total mRNA number provides the number of cytoplasmic mRNA spots. Our findings reveal that approximately 92% of Mef2 mRNA is present in the cytoplasm, while the remaining 8% is associated with muscle nuclei, as shown in Figure 3E,F.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65713/65713fig1v2.jpg" /> 
Figure 1: Procedure to dissect and prepare wing discs and IFM samples for smFISH. (A) L3 staged larvae. (B)&#160;Dissected and inverted anterior end of the larva. Arrows indicate the wing discs. (C) Isolated wing disc. (D) Dissected adult thoraxes. (E) Adult thorax oriented on a doublesided tape prior to bisection (dotted line). (F) Adult hemithorax. (G) Isolated IFMs. (H) Workflow of the smFISH protocol. Abbreviations: IFMs = indirect flight muscles; smFISH = single-molecule RNA fluorescence in situ hybridization; EtOH = ethanol. <a href="https://www.jove.com/files/ftp_upload/65713/65713fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65713/65713fig2v2.jpg" /> 
Figure 2: Representative images of Mef2 and zfh1 smFISH in wing discs and adult IFMs. (A,B) Mef2 and zfh1 mRNAs (purple) are detected uniformly in AMPs and co-localize with (A,A&#39;) Mef2 and (B,B&#39;) Zfh1 proteins (green), respectively. (A&#39;&#39;,B&#39;&#39;)&#160;Higher magnification of the AMPs. (C)&#160;Mef2 mRNAs and Mef2 protein are detected in adult IFMs. (C&#39;) Higher magnification of the boxed region in C. (D)&#160;Zfh1-Gal4 &#62;UAS-mCD8GFP expression (green) labels adult MuSCs. zfh1 RNA is detected in adult MuSCs. (D&#39;) Higher magnification of the boxed region in D. Zfh1 transcription start sites and single mRNAs are indicated by arrows and arrowheads, respectively. DAPI staining (blue). Scale bars = 50 &#181;m (A,B,A&#39;,B&#39;), 10 &#181;m (A&#34;,B&#34;,C,D), 5 &#181;m (C&#39;,D&#39;). Abbreviations: IFMs = indirect flight muscles; smFISH = single-molecule RNA fluorescence in situ hybridization; AMPs = adult muscle progenitors; MuSCs = muscle stem cells; DAPI = 4&#39;,6-diamidino-2-phenylindole; GFP = green fluorescent protein. <a href="https://www.jove.com/files/ftp_upload/65713/65713fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65713/65713fig3v2.jpg" /> 
Figure 3: Quantification of Mef2 transcription and cytoplasmic mRNA accumulation by smFISH. (A) Distribution of Mef2 transcripts (purple) and Mef2 protein (green) in wildtype adult IFMs. (A&#39;) Higher magnification of boxed region in A. Transcription start sites and single mRNAs are indicated by arrows and arrowheads, respectively. (B,C) Automatic Mef2 spots finding and nuclear segmentation. Cytoplasmic and nuclear Mef2 mRNAs are indicated in green and magenta, respectively. (B&#39;,C&#39;) Higher magnifications of boxed regions in B and C. Asterisks indicate spots that are not counted by the macro. (D) Example of Mef2 spot quantification in AMPs and adult IFMs, relative to the total number of nuclei. (p = 0,0785, Student&#39;s t-test, wing discs (n = 4), IFMs (n = 11)). (E) Example of quantification of Mef2 spot distribution in adult IFMs. (*** p&#60; 0.0007, Student&#39;s t-test, n = 5). (F) Percentage of Mef2 mRNA spots detected inside and outside the nuclei (n = 5). Scale bars = 10 &#181;m (A,A&#39;). Abbreviations: IFMs = indirect flight muscles; smFISH = single-molecule RNA fluorescence in situ hybridization; AMPs = adult muscle progenitors. <a href="https://www.jove.com/files/ftp_upload/65713/65713fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental File 1: Macro used for postprocessing. <a href="https://www.jove.com/files/ftp_upload/65713/Le Roux et al-Macro.txt" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Investigating mRNA Spatial Distribution in Drosophila Muscle Tissue at JoVE.com

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.

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 ...

studying-muscle-transcriptional-dynamics-at-single-molecule-scales

Research

JoVE Journal

Developmental Biology

1.4K Views.  UMR6290 CNRS - Université de Rennes. In ons team bestudeerden we de cellulaire en moleculaire mechanismen die de ontwikkeling en het herstel van spieren regelen. Het is zelfs bekend dat de speciale verdeling van mRNA-moleculen verschillende cellulaire processen reguleert. En hier gebruiken we de Drosophila als model om te onderzoeken hoe de mRNA's vooral verdeeld zijn in het spierweefsel van de vlieg.Het is bewezen dat genexpressiedynamiek verschillende spierbiologische processen reguleert. De komst van high-throughput si...

Video: Auteur in de schijnwerpers: onderzoek naar ruimtelijke mRNA-verdeling in Drosophila-spierweefsel 

Improved Protocol for Chromatin Immunoprecipitation from Mouse Skeletal Muscle

We describe an efficient and reproducible protocol for the preparation of chromatin from adult mouse skeletal muscle, a physically resistant tissue with a high content of structural proteins. Dissected limb muscles from adult mice are physically disrupted by mechanical homogenisation, or a combination of mincing and douncing, in a hypotonic buffer before formaldehyde fixation of the cell lysate. The fixed nuclei are purified by further cycles of mechanical homogenisation or douncing and sequential filtrations to remove cell debris. The purified nuclei can be sonicated immediately or at a later stage after freezing. The chromatin can be efficiently sonicated and is suitable for chromatin immunoprecipitation experiments, as illustrated by the profiles obtained for transcription factors, RNA polymerase II, and covalent histone modifications. The binding events detected using chromatin prepared by this protocol are predominantly those taking place in the muscle fiber nuclei despite the presence of chromatin from other fiber-associated satellite and endothelial cells. This protocol is therefore adapted to study gene regulation in the adult mouse skeletal muscle.

A novel protocol for the preparation of chromatin from adult mouse skeletal muscle adapted to the study of gene regulation in muscle fibers by chromatin immunoprecipitation is presented.

We describe an efficient and reproducible protocol for the preparation of chromatin from adult mouse skeletal muscle, a physically resistant tissue with a ...

Biochemistry

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.

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.

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

Genetics

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from animals to humans. Therefore, the powerful Drosophila model system can be used to study concepts of muscle-tendon development that can also be applied to human muscle biology. Here, we describe in detail how morphogenesis of the adult muscle-tendon system can be easily imaged in living, developing Drosophila pupae. Hence, the method allows investigating proteins, cells and tissues in their physiological environment. In addition to a step-by-step protocol with helpful tips, we provide a comprehensive overview of fluorescently tagged marker proteins that are suitable for studying the muscle-tendon system. To highlight the versatile applications of the protocol, we show example movies ranging from visualization of long-term morphogenetic events &#8211; occurring on the time scale of hours and days &#8211; to visualization of short-term dynamic processes like muscle twitching occurring on time scale of seconds. Taken together, this protocol should enable the reader to design and perform live-imaging experiments for investigating muscle-tendon morphogenesis in the intact organism.

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Here, we present an easy-to-use and versatile method to perform live imaging of developmental processes in general and muscle-tendon morphogenesis in particular in living Drosophila pupae.

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

Single Nucleotide Polymorphism-sensitive FISH Detection of Locus-specific Ribosomal RNA Transcription in Drosophila melanogaster

In order to ensure sufficient ribosomal RNA (rRNA) is transcribed for ribosome function, genomes contain hundreds of tandem duplications of the sequences that encode rRNAs, composing regions called ribosomal DNA (rDNA) loci. Many organisms' genomes contain more than one rDNA locus distributed across different chromosomes, and rRNA may be transcribed from multiple rDNA loci or only a single locus. Changes in rRNA transcription sources often indicate ribosomal distress. However, the homogeneity of rRNA obstructs distinguishing if rRNA is transcribed from multiple or a single rDNA locus. Here, we describe a method that uses single nucleotide polymorphism-sensitive fluorescence in situ hybridization (SNP-FISH) to distinguish rRNA transcription between Drosophila melanogaster rDNA loci on the X and Y chromosomes. This method leverages locus-specific rRNA variants to enable easy detection of the source of rRNA transcription with single-cell resolution. This assay can be applied to any Drosophila cell type and may be adapted for use in other systems.

The protocol describes single nucleotide polymorphisms-sensitive fluorescence in situ hybridization (SNP-FISH) method to distinguish ribosomal RNA transcripts derived from the X or Y chromosome ribosomal DNA locus in Drosophila melanogaster.

In order to ensure sufficient ribosomal RNA (rRNA) is transcribed for ribosome function, genomes contain hundreds of tandem duplications of the sequences ...

Chromatin Immunoprecipitation (ChIP) using Drosophila tissue

Epigenetics remains a rapidly developing field that studies how the chromatin state contributes to differential gene expression in distinct cell types at different developmental stages. Epigenetic regulation contributes to a broad spectrum of biological processes, including cellular differentiation during embryonic development and homeostasis in adulthood. A critical strategy in epigenetic studies is to examine how various histone modifications and chromatin factors regulate gene expression. To address this, Chromatin Immunoprecipitation (ChIP) is used widely to obtain a snapshot of the association of particular factors with DNA in the cells of interest.
ChIP technique commonly uses cultured cells as starting material, which can be obtained in abundance and homogeneity to generate reproducible data. However, there are several caveats: First, the environment to grow cells in Petri dish is different from that in vivo, thus may not reflect the endogenous chromatin state of cells in a living organism. Second, not all types of cells can be cultured ex vivo. There are only a limited number of cell lines, from which people can obtain enough material for ChIP assay.
Here we describe a method to do ChIP experiment using Drosophila tissues. The starting material is dissected tissue from a living animal, thus can accurately reflect the endogenous chromatin state. The adaptability of this method with many different types of tissue will allow researchers to address a lot more biologically relevant questions regarding epigenetic regulation in vivo1, 2. Combining this method with high-throughput sequencing (ChIP-seq) will further allow researchers to obtain an epigenomic landscape.

Chromatin Immunoprecipitation ChIP using Drosophila tissue

Recently high-throughput sequencing technology has greatly increased sensitivity of Chromatin Immunoprecipitation (ChIP) experiment and prompted its application using purified cells or dissected tissue. Here we delineate a method to use ChIP technique with Drosophila tissue, which can address the endogenous chromatin state in a well-characterized biological system.

Epigenetics remains a rapidly developing field that studies how the chromatin state contributes to differential gene expression in distinct cell types at ...

Biology

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Neuroscience

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. Furthermore, despite progress in understanding postsynaptic MTs, much less is known about the contributions of presynaptic MTs to neuronal morphogenesis. In particular, studies of in vivo MT dynamics in Drosophila sensory dendrites yielded significant insights into polymer-level behavior. However, the technical and analytical challenges associated with live imaging of the fly neuromuscular junction (NMJ) have limited comparable studies of presynaptic MT dynamics. Moreover, while there are many highly effective software strategies for automated analysis of MT dynamics in vitro and ex vivo, in vivo data often necessitate significant operator input or entirely manual analysis due to inherently inferior signal-to-noise ratio in images and complex cellular morphology. &#160;To address this, this study optimized a new software platform for automated and unbiased in vivo particle detection. Multiparametric analysis of live time-lapse confocal images of EB1-GFP labeled MTs was performed in both dendrites and the NMJ of Drosophila larvae and found striking differences in MT behaviors. MT dynamics were furthermore analyzed following knockdown of the MT-associated protein (MAP) dTACC, a key regulator of Drosophila synapse development, and identified statistically significant changes in MT dynamics compared to wild type. These results demonstrate that this novel strategy for the automated multiparametric analysis of both pre- and postsynaptic MT dynamics at the polymer-level significantly reduces human-in-the-loop criteria. The study furthermore shows the utility of this method in detecting distinct MT behaviors upon dTACC-knockdown, indicating a possible future application for functional screens of factors that regulate MT dynamics in vivo. Future applications of this method may also focus on elucidating cell type and/or compartment-specific MT behaviors, and multicolor correlative imaging of EB1-GFP with other cellular and subcellular markers of interest.&#160;

This study presents a noninvasive intravital neuronal imaging strategy combined with a new software strategy to achieve automated, unbiased tracking and analysis of in vivo microtubule (MT) plus-end dynamics in the sensory dendrites and the neuromuscular junctions of Drosophila.

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. ...

TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos

Measuring levels of mRNAs in the process of translation in individual cells provides information on the proteins involved in cellular functions at a given point in time. The protocol dubbed Translating Ribosome Affinity Purification (TRAP) is able to capture this mRNA translation process in a cell-type-specific manner. Based on the affinity purification of polysomes carrying a tagged ribosomal subunit, TRAP can be applied to translatome analyses in individual cells, making it possible to compare cell types during the course of developmental processes or to track disease development progress and the impact of potential therapies at molecular level. Here we report an optimized version of the TRAP protocol, called TRAP-rc (rare cells), dedicated to identifying engaged-in-translation RNAs from rare cell populations. TRAP-rc was validated using the Gal4/UAS targeting system in a restricted population of muscle cells in Drosophila embryos. This novel protocol allows the recovery of cell-type-specific RNA in sufficient quantities for global gene expression analytics such as microarrays or RNA-seq. The robustness of the protocol and the large collections of Gal4 drivers make TRAP-rc a highly versatile approach with potential applications in cell-specific genome-wide studies.

TRAP-rc, Translating Ribosome Affinity Purification from Rare Cell Populations of Drosophila Embryos

Translating Ribosome Affinity Purification (TRAP) is able to capture cell-type-specific translation of mRNA. Here we report the first TRAP protocol dedicated to isolation of mRNA in rare cell populations of Drosophila embryos.

Measuring levels of mRNAs in the process of translation in individual cells provides information on the proteins involved in cellular functions at a given ...

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 ...

Studying Muscle Transcriptional Dynamics at Single-molecule Scales in Drosophila

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