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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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mef2+and+the+skeletal+muscle+differentiation+program.">Mef2 and the skeletal muscle differentiation program.</a> Seminars in Cell and Developmental Biology. 72, 33-44 (2017).</li><li>Siles, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ZEB1+imposes+a+temporary+stage-dependent+inhibition+of+muscle+gene+expression+and+differentiation+via+CtBP-mediated+transcriptional+repression.">ZEB1 imposes a temporary stage-dependent inhibition of muscle gene expression and differentiation via CtBP-mediated transcriptional repression.</a> Molecular and Cellular Biology. 33 (7), 1368-1382 (2013).</li><li>Weitkunat, M., Schnorrer, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+to+study+Drosophila+muscle+biology.">A guide to study Drosophila muscle biology.</a> Methods. 68 (1), 2-14 (2014).</li><li>Boukhatmi, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila,+an+integrative+model+to+study+the+features+of+musclestem+cells+in+development+and+regeneration.">Drosophila, an integrative model to study the features of musclestem cells in development and regeneration.</a> Cells. 10 (8), 2112 (2021).</li><li>Bothe, I., Baylies, M. 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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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160;</td>
			<td>Tesa&#160;</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&#160;</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

856 Views.  UMR6290 CNRS - Université de Rennes. 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. 

Video: Author Spotlight: Investigating mRNA Spatial Distribution in Drosophila Muscle Tissue 

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Imaging Calcium in Drosophila at Egg Activation

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. One aspect of egg activation, conserved across all organisms examined, is a change in the intracellular concentration of calcium (Ca2+) often termed a 'Ca2+ wave'. While the speed and number of oscillations of the Ca2+ wave varies between species, the change in intracellular Ca2+ is key in bringing about essential events for embryonic development. These changes include resumption of the cell cycle, mRNA regulation, cortical granule exocytosis, and rearrangement of the cytoskeleton.
In the mature Drosophila egg, activation occurs in the female oviduct prior to fertilization, initiating a series of Ca2+-dependent events. Here we present a protocol for imaging the Ca2+ wave in Drosophila. This approach provides a manipulable model system to interrogate the mechanism of the Ca2+ wave and the downstream changes associated with it.

Imaging Calcium in Drosophila at Egg Activation

This article describes an adaptable ex vivo protocol for visualizing Ca2+ during egg activation in Drosophila.

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. ...

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

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 ...

A Cell-based Assay to Investigate Non-muscle Myosin II Contractility via the Folded-gastrulation Signaling Pathway in Drosophila S2R+ Cells

We have developed a cell-based assay using Drosophila cells that recapitulates apical constriction initiated by folded gastrulation (Fog), a secreted epithelial morphogen. In this assay, Fog is used as an agonist to activate Rho through a signaling cascade that includes a G-protein-coupled receptor (Mist), a G&#945;12/13 protein (Concertina/Cta), and a PDZ-domain-containing guanine nucleotide exchange factor (RhoGEF2). Fog signaling results in the rapid and dramatic reorganization of the actin cytoskeleton to form a contractile purse string. Soluble Fog is collected from a stable cell line and applied ectopically to S2R+ cells, leading to morphological changes like apical constriction, a process observed during developmental processes such as gastrulation. This assay is amenable to high-throughput screening and, using RNAi, can facilitate the identification of additional genes involved in this pathway.

A Cell-based Assay to Investigate Non-muscle Myosin II Contractility via the Folded-gastrulation Signaling Pathway in Drosophila S2R+ Cells

Here we describe a contractility assay using Drosophila S2R+ cells. The application of an exogenous ligand, folded gastrulation (Fog), leads to the activation of the Fog signaling pathway and cellular contractility. This assay can be used to investigate the regulation of cellular contractility proteins in the Fog signaling pathway.

We have developed a cell-based assay using Drosophila cells that recapitulates apical constriction initiated by folded gastrulation (Fog), a secreted ...

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...

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 ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Drosophila cell lines are important reagents for both fundamental and biomedical research. This article provides protocols for thawing, subculturing, and the cryopreservation of commonly used Drosophila cell lines to assist researchers in incorporating the use of these reagents in their research.

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the ...

X-ray Diffraction of Intact Murine Skeletal Muscle as a Tool for Studying the Structural Basis of Muscle Disease

Transgenic mouse models have been important tools for studying the relationship of genotype to phenotype for human diseases including those of skeletal muscle. Mouse skeletal muscle has been shown to produce high quality X-ray diffraction patterns on third generation synchrotron beamlines providing an opportunity to link changes at the level of the genotype to functional phenotypes in health and disease by determining the structural consequences of genetic changes. We present detailed protocols for preparation of specimens, collecting the X-ray patterns and extracting relevant structural parameters from the X-ray patterns that may help guide experimenters wishing to perform such experiments for themselves.

We present detailed protocols for performing small-angle X-ray diffraction experiments using intact mouse skeletal muscles. With the wide availability of transgenic mouse models for human diseases, this experimental platform can form a useful test bed for elucidating the structural basis of genetic muscle diseases

Transgenic mouse models have been important tools for studying the relationship of genotype to phenotype for human diseases including those of skeletal ...