ai_agent

We thank Manuela Weitkunat for the acquisition of Movie S3. We are grateful to Reinhard F&#228;ssler for generous support. This work was supported by the EMBO Young Investigator Program (F.S.), the European Research Council under the European Union&#39;s Seventh Framework Programme (FP/2007-2013)/ERC Grant 310939 (F.S.), the Max Planck Society (S.B.L., F.S.), the Centre National de la Recherche Scientifique (CNRS) (F.S.), the excellence initiative Aix-Marseille University AMIDEX (F.S.), the LabEX-INFORM (F.S.) and the Boehringer Ingelheim Fonds (S.B.L.).

1. Stage Pupae
<ol>
	<li>Set up a cross with 20 - 40 virgin females and about 20 males per bottle of fly food (Figure 1A). Alternatively, if stocks will be used, flip each stock into new bottles. To get enough pupae, consider setting up two bottles per genotype.</li>
	<li>Incubate the bottles at 25 °C (or 27 °C according to the experimental design) for five to six days (Figure 1B). 
	NOTE: Keep flipping the flies every two to three days to avoid overcrowding and to ensure a continuous supply of pupae.</li>
	<li>Use a wet brush to collect white prepupae from the walls of the bottles and transfer them to the glass slides (Figure 1C). Time the staging such that the pupae reach the desired age when starting live imaging under the microscope. 
	NOTE: These prepupae look like older pupae in terms of their shape, but they are still white like larvae. The onset of the prepupal stage is defined as 0 h after puparium formation (0 h APF).</li>
	<li>Remove clumps of fly food sticking to the pupae with a wet brush and orient the ventral side of the pupae towards the glass slide using a stereomicroscope (Figure 1D). Discard pupae that are still moving (too young) or that have started to turn brown (too old). This ensures that all pupae have the same age (0 - 1 h APF).</li>
	<li>Label the slides with genotype, date and time of collection. Transfer the glass slides to one petri dish each and add a wet tissue to prevent the pupae from drying out (Figure 1E).</li>
	<li>Store the pupae in an incubator at 25 °C or 27 °C until they reach the desired age. Continue with the next steps 1 h before imaging, so that the pupae have the desired age when starting live imaging (step 3), for example at 13 h APF for the movie shown in Figure 2A-D. 
	NOTE: The earliest feasible starting time point is after head-eversion of the pupae occurring at 8 - 10 h APF at 27 °C.</li>
</ol>
2. Prepare Pupae for Imaging
<ol>
	<li>Ensure that the pupae now stick to the glass slide just like they naturally stick to the walls of the bottles because of residual fly food. No extra glue is required. If the pupae do not stick well enough because of high humidity, open the petri dish for a few min to let them dry. Alternatively, transfer them to double-sided tape on a glass slide, but usually, this is not necessary.</li>
	<li>Open window in the pupal case for imaging of indirect flight muscles (Figure 1F; Skip to section 2.3 for imaging of abdominal muscles.)
	<ol>
		<li>Orient the pupae sticking to the glass slide with the anterior facing away from the researcher under a stereomicroscope. Adjust the zoom to 2X.</li>
		<li>Using one end of biology grade #5 forceps, gently poke a hole into the pupal case dorsal to the wing where the abdomen ends, and the thorax begins.</li>
		<li>To slice open the pupal case, gently move the forceps to the anterior end of the thorax along the wing but avoid damaging the vulnerable wing tissue. 
		NOTE: If fluid leaks from the pupa, it is damaged; discard it and start over with a different pupa.</li>
		<li>Lift up the section of the pupal case above the thorax with the forceps and then cut this section off with fine, sharp scissors on the opposite side of the dorsal midline.</li>
	</ol>
	</li>
	<li>Open window in pupal case for imaging of abdominal muscles (Figure 1G)
	<ol>
		<li>Orient the pupae sticking to the glass slide with the anterior facing towards the researcher under a stereomicroscope. Adjust the zoom to 2X.</li>
		<li>Using one end of biology grade #5 forceps, gently poke a hole into the pupal case dorsal to the wing where the thorax ends, and the abdomen begins.</li>
		<li>To slice open the pupal case, gently move the forceps towards the posterior end of the abdomen. 
		NOTE: If fluid leaks from the pupa, it is damaged; discard it and start over with a different pupa.</li>
		<li>Lift up the section of the pupal case above the abdomen with the forceps and then cut this section off with fine, sharp scissors on the opposite side of the dorsal midline.</li>
	</ol>
	</li>
	<li>Mount Pupae (Figure 1H)
	<ol>
		<li>Use a wet brush to transfer up to five pupae to a plastic slide with a groove (custom-built, reusable, see discussion and list of materials). Add one or two spacer coverslips to each side depending on the depth of the groove and the thickness of the pupae. Put a small drop of water underneath each spacer coverslip to make it stick to the slide and leave some space between the groove and the spacer coverslips on each side. 
		NOTE: The spacer coverslips can also be permanently attached to the slide with super glue, in advance.</li>
		<li>Orient the pupae such that the opening in the pupal case faces upwards using the brush. Ensure careful positioning of the pupae at the correct angle for good imaging quality. Optimize the angle for each tissue and developmental time point. 
		NOTE: A small amount of water in the groove makes positioning of the pupae easier. Drain excess water with the brush afterward to avoid drowning.</li>
		<li>Hold a coverslip (18 x 18 mm) above the pupae without touching them. Place small droplets (about 0.5 µL) of 50% glycerol solution above each pupal case opening onto the coverslip using a 20 µL pipette. 
		NOTE: This procedure ensures that the droplets have the same spacing as the pupae.</li>
		<li>Turn the coverslip over and position it above the pupae by hand or using forceps (standard #5) while resting one side of the coverslip on the spacer coverslips next to the pupae. Next, gently drop the coverslip onto the pupae. Ensure that the pupal case openings are properly covered with 50% glycerol for good imaging quality and to prevent the pupae from drying out.</li>
		<li>Fold over one end of a piece of adhesive tape and grab it with forceps (standard #5) at that end. Gently place the tape such that it covers one edge of the top coverslip, the spacer coverslip(s), and the plastic slide on one side. Do not press down the tape, yet. First, turn the slide around and repeat for the other side.</li>
		<li>Use both index fingers to press down the tape simultaneously on both sides. This procedure ensures that the pupae are not displaced.</li>
		<li>Check whether the pupae are well in contact with the coverslip. For optimal imaging quality the pupae should be gently squeezed underneath the coverslip. Adjust the number of spacer coverslips if necessary.</li>
		<li>Label the slide by writing on the sticky tape.</li>
	</ol>
	</li>
</ol>
3. Live Imaging of Pupae
<ol>
	<li>The mounting method allows imaging of the pupae both on inverted microscopes (Figure 1I) and on upright microscopes (Figure 1J). Especially suitable for live imaging are scanning confocal microscopes, two-photon microscopes and spinning disc confocal microscopes irrespective of whether they are inverted or upright. For long-term movies, use a temperature-controlled stage if available.</li>
	<li>Using the ocular and a UV lamp or transmission light, locate a pupa and focus inside the pupa.</li>
	<li>Using the camera, find the desired structure and adjust the zoom level.</li>
	<li>For long-term movies, define a z-stack that encompasses the structure of interest well (for example, the indirect flight muscles, their attachment sites, the tendon cells or a combination the aforementioned) and choose a time interval. Consider doing averaging for better image quality. For high-speed, short-term movies, choose a single z-plane to achieve a high frame rate.</li>
	<li>Adjust the laser power to give the best possible signal but little saturation. However, too high laser power can damage the pupa over time.</li>
	<li>Start imaging and return from time to time to check the movie and readjust the z-stack positioning if necessary.</li>
</ol>
<img alt="Drosophila pupae preparation and imaging process for muscle study; includes microscope setup." class="xfigimg" src="/files/ftp_upload/57312/57312fig1.jpg" data-altupdatedat="1762215414000" data-alt="Figure 1"> 
Figure 1: Workflow for live imaging of muscle-tendon morphogenesis in Drosophila pupae. See protocol for details. <a href="//cloudfront.jove.com/files/ftp_upload/57312/57312fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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The authors have nothing to disclose.

The presented protocol describes how to image muscle-tendon morphogenesis in living Drosophila pupae using a variety of fluorescently tagged proteins. This in vivo imaging strategy can be used to study developmental processes in their natural environment of the entire organism.
It is crucial for a successful experiment to find the correct developmental time point to analyze. For example, dorsolongitudinal indirect flight muscles initiate attachment to their tendon targets at &#8776;16 h APF23 while abdominal muscles develop later and attach on both ends only between 30 and 40 h APF26. Consequently, previously published literature should be used to find the right time points of development to analyze or, if the tissue or structure of interest has not been studied in detail before, the overall development has to be characterized first.
For mounting pupae successfully on the custom-built plastic slides, it is important that the grooves have suitable dimensions: The grooves need to be 1.0 - 1.5 mm wide and 0.3 - 0.4 mm deep. This depth allows adjusting the precise distance to the top coverslip with spacer coverslips as needed. However, at least one spacer coverslip should be used to avoid draining the 50% glycerol away from the sample by capillary forces. The correct positioning of the pupae in the groove requires some experience and should be optimized such that the structure of interest is as close as possible to the coverslip.
If a large number of pupae is supposed to be imaged in one microscope session, they can all be mounted beforehand and then stored in an incubator until imaging to ensure proper developmental timing. The pupae should survive the entire procedure and also at least try to eclose if kept on the slide after imaging. The survival rate can be used as a readout to check whether the imaging conditions harm the pupae.
The imaging settings should be chosen carefully according to the experimental requirements. For short-term movies, a high frame rate versus a high signal-to-noise ratio needs to be balanced, while relatively high laser power can be used without damaging the pupae too much. However, for long-term movies, the laser power has to be kept at a moderate level and the pupae should not be imaged continuously but rather at certain time points, for example, every 20 min. To ensure that the structure of interest does not move out of the field of view, it might be necessary to readjust the positioning of the z-stack between time points. To our knowledge, the opening of the pupal case per se does not affect developmental timing. However, a temperature-controlled stage should be used for long-term movies to ensure proper developmental timing. Keeping these considerations in mind, highly informative movies can be acquired.
The presented protocol can be used to visualize not only muscle-tendon morphogenesis but also other developing tissues, for example, the wing epithelium29. Only three modifications to this protocol are required: (1) opening of the pupal case above the wing instead of the thorax or abdomen, (2) positioning of pupae with the wing towards the top coverslip, and (3) the use of different fluorescent marker proteins. With the advancement of the CRISPR/Cas9-technology, more and more endogenously tagged fluorescent proteins will be available, because it has become more straightforward to target endogenous loci in Drosophila30,31,32. In the future, this will allow elucidating the dynamics of numerous proteins, cells and entire tissues in their physiological environment in detail.

The muscle-tendon apparatus allows animals including humans to move their body parts. The molecular building blocks of the muscle-tendon system are highly conserved. Therefore, concepts of muscle-tendon development relevant for human muscle biology, for example muscle morphogenesis, muscle-tendon attachment and myofibril self-organization, can be studied using Drosophila melanogaster as an easily accessible model system. The Drosophila pupal system has several experimental advantages. First, at the pupal stage &#8211; when the adult muscles are formed &#8211; the organism is sessile and therefore easy to image on a microscope over a period of hours or even days. Second, many muscles form close enough underneath the pupal surface so that they can be imaged inside the intact, partially translucent organism. Third, the muscles can be investigated in their natural environment, where they are connected to the forming exoskeleton via tendon cells and tissue tension is built up. This is not possible in muscle cell culture systems. And finally, a plethora of genetic tools is available in Drosophila. Among these are many fluorescently tagged markers that allow labeling of specific cell types or subcellular structures for imaging in vivo.
Table 1 summarizes the most important markers used for studying muscle-tendon morphogenesis. It includes markers overexpressed using the GAL4-UAS-system1 and endogenously tagged protein markers2,3,4. The advantage of the GAL4-UAS-system is that the markers are generally expressed at high levels, resulting in a strong signal that can easily be imaged in whole-mount pupae. In addition, tissue specificity can be achieved by choosing GAL4 drivers carefully. The advantage of fusion proteins expressed under endogenous control is that the dynamics of the respective proteins can be studied in vivo, while they can also be used as markers for different cell types or specific subcellular structures, for example, &#946;PS-Integrin-GFP for muscle attachment sites. Together, these markers provide high flexibility in experimental design and choice of research problems that can be solved now and in the future.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Labeled structure</td>
			<td colspan="2">Marker</td>
			<td colspan="2">Expression and localization</td>
			<td colspan="2">Class</td>
			<td colspan="2">Stock number</td>
			<td colspan="2">Comment </td>
			<td>Ref.</td>
		</tr>
		<tr>
			<td colspan="2" rowspan="15">Muscle</td>
			<td colspan="2">Mef2-GAL4</td>
			<td colspan="2">all myoblasts and all muscles at all stages</td>
			<td colspan="2">GAL4 line</td>
			<td colspan="2">BL 27390</td>
			<td colspan="2"></td>
			<td>5</td>
		</tr>
		<tr>
			<td colspan="2">1151-GAL4</td>
			<td colspan="2">adult muscle precursors and early myotubes until &#8776;24 h APF</td>
			<td colspan="2">GAL4 line, enhancer trap</td>
			<td colspan="2">-</td>
			<td colspan="2"></td>
			<td>6</td>
		</tr>
		<tr>
			<td colspan="2">Act79B-GAL4</td>
			<td colspan="2">jump muscle upon differentiation</td>
			<td colspan="2">GAL4 line</td>
			<td colspan="2">-</td>
			<td colspan="2"></td>
			<td>7</td>
		</tr>
		<tr>
			<td colspan="2">Act88F-GAL4</td>
			<td colspan="2">indirect flight muscles starting &#8776;14 h APF</td>
			<td colspan="2">GAL4 line</td>
			<td colspan="2">-</td>
			<td colspan="2"></td>
			<td>7</td>
		</tr>
		<tr>
			<td colspan="2">Act88F-Cameleon 3.1</td>
			<td colspan="2">indirect flight muscles starting &#8776;14 h APF</td>
			<td colspan="2">Act88F enhancer/ promoter driving Cameleon 3.1</td>
			<td colspan="2">-</td>
			<td colspan="2">Ca2+ indicator</td>
			<td>8</td>
		</tr>
		<tr>
			<td colspan="2">Act88F-GFP</td>
			<td colspan="2">indirect flight muscles starting &#8776;14 h APF</td>
			<td colspan="2">GFP-fusion (fly TransgeneOme line)</td>
			<td colspan="2">fTRG78 and fTRG10028</td>
			<td colspan="2"></td>
			<td>4</td>
		</tr>
		<tr>
			<td colspan="2">Him-nls-GFP</td>
			<td colspan="2">adult muscle precursors, nuclear, until &#8776;24 h APF in indirect flight muscles</td>
			<td colspan="2">enhancer/promoter with nls-GFP reporter</td>
			<td colspan="2">-</td>
			<td colspan="2">1.5 kb enhancer fragment</td>
			<td>9</td>
		</tr>
		<tr>
			<td colspan="2">Mhc-Tau-GFP</td>
			<td colspan="2">microtubules in DLM templates and in differentiating muscles</td>
			<td colspan="2">enhancer/promoter with Tau-GFP reporter</td>
			<td colspan="2">BL 53739</td>
			<td colspan="2"></td>
			<td>10</td>
		</tr>
		<tr>
			<td colspan="2">&#946;Tub60D-GFP</td>
			<td colspan="2">microtubules in myotubes (e.g. in indirect flight muscles from &#8776;14 h AFP, strongly decreasing after &#8776;48 h APF)</td>
			<td colspan="2">GFP-fusion (fly TransgeneOme line)</td>
			<td colspan="2">fTRG958</td>
			<td colspan="2"></td>
			<td>4</td>
		</tr>
		<tr>
			<td colspan="2">Mhc-GFP (weeP26)</td>
			<td colspan="2">sarcomeres (thick filament) in all body muscles (e.g. in indirect flight muscles starting from &#8776;30 h APF)</td>
			<td colspan="2">GFP-trap</td>
			<td colspan="2">-</td>
			<td colspan="2">use heterozygous, labels an isoform subset</td>
			<td>11</td>
		</tr>
		<tr>
			<td colspan="2">Sls-GFP</td>
			<td colspan="2">sarcomeres (Z-disc) in all body muscles (e.g. in indirect flight muscles starting from &#8776;30 h APF)</td>
			<td colspan="2">GFP-trap (FlyTrap line)</td>
			<td colspan="2">-</td>
			<td colspan="2">G53, use heterozygous</td>
			<td>2</td>
		</tr>
		<tr>
			<td colspan="2">Zasp66-GFP</td>
			<td colspan="2">Z-disc in all body muscles</td>
			<td colspan="2">GFP-trap (FlyTrap line)</td>
			<td colspan="2">BL 6824</td>
			<td colspan="2">ZCL0663</td>
			<td>2,12</td>
		</tr>
		<tr>
			<td colspan="2">Zasp52-GFP</td>
			<td colspan="2">Z-disc in all body muscles</td>
			<td colspan="2">GFP-trap (FlyTrap line)</td>
			<td colspan="2">BL 6838</td>
			<td colspan="2">G00189</td>
			<td>2,12</td>
		</tr>
		<tr>
			<td colspan="2">Hts-GFP</td>
			<td colspan="2">actin binding; expressed in epithelium, myoblasts and myotubes</td>
			<td colspan="2">GFP-fusion (fly TransgeneOme line)</td>
			<td colspan="2">fTRG585</td>
			<td colspan="2"></td>
			<td>4</td>
		</tr>
		<tr>
			<td colspan="2">Dlg1-GFP</td>
			<td colspan="2">epithelial cell junctions, myoblasts and membranes in muscles at all stages</td>
			<td colspan="2">GFP-fusion (fly TransgeneOme line)</td>
			<td colspan="2">fTRG502</td>
			<td colspan="2"></td>
			<td>4</td>
		</tr>
		<tr>
			<td colspan="2" rowspan="5">Muscle attachment site</td>
			<td colspan="2">&#946;PS-Integrin-GFP</td>
			<td colspan="2" fo:page-break-inside="avoid">muscle attachment sites (e.g., starting &#8776;18 h AFP in indirect flight muscles)</td>
			<td colspan="2">GFP-knock-in</td>
			<td colspan="2">-</td>
			<td colspan="2"></td>
			<td>13</td>
		</tr>
		<tr>
			<td colspan="2">Talin-GFP and -mCherry</td>
			<td colspan="2">muscle attachment sites (e.g., starting &#8776;18 h AFP in indirect flight muscles)</td>
			<td colspan="2">GFP-trap (MiMIC line)</td>
			<td colspan="2">-</td>
			<td colspan="2"></td>
			<td>3</td>
		</tr>
		<tr>
			<td colspan="2">Talin-GFP</td>
			<td colspan="2">muscle attachment sites (e.g., starting &#8776;18 h AFP in indirect flight muscles)</td>
			<td colspan="2">GFP-fusion (fly TransgeneOme line)</td>
			<td colspan="2">fTRG587</td>
			<td colspan="2"></td>
			<td>4</td>
		</tr>
		<tr>
			<td colspan="2">Ilk-GFP</td>
			<td colspan="2">muscle attachment sites (e.g., starting &#8776;18 h AFP in indirect flight muscles)</td>
			<td colspan="2">GFP-trap (FlyTrap line)</td>
			<td colspan="2">Kyoto 110951 (ZCL3111)</td>
			<td colspan="2">ZCL3111, ZCL3192</td>
			<td>2</td>
		</tr>
		<tr>
			<td colspan="2">Vinc-GFP and -RFP</td>
			<td colspan="2">muscle attachment sites (e.g., starting &#8776;18 h AFP in indirect flight muscles)</td>
			<td colspan="2">GFP-fusion (transgene)</td>
			<td colspan="2">-</td>
			<td colspan="2"></td>
			<td>13</td>
		</tr>
		<tr>
			<td colspan="2">Tendon</td>
			<td colspan="2">sr-GAL4</td>
			<td colspan="2">thorax tendon cells, throughout pupal stage</td>
			<td colspan="2">GAL4 line, enhancer trap</td>
			<td colspan="2">BL 26663</td>
			<td colspan="2">homozygous lethal</td>
			<td>14</td>
		</tr>
		<tr>
			<td colspan="2">Muscle and Tendon</td>
			<td colspan="2">Duf-GAL4</td>
			<td colspan="2">muscles and epithelia, early onset</td>
			<td colspan="2">GAL4 line</td>
			<td colspan="2">BL 66682</td>
			<td colspan="2">kirre-rP298, founder cell marker</td>
			<td>15</td>
		</tr>
		<tr>
			<td colspan="2" rowspan="7">UAS-reporters</td>
			<td colspan="2">UAS-GFP-Gma</td>
			<td colspan="2">actin binding</td>
			<td colspan="2">UAS line</td>
			<td colspan="2">BL 31776</td>
			<td colspan="2">actin binding domain of Moesin fused to GFP</td>
			<td>16</td>
		</tr>
		<tr>
			<td colspan="2">UAS-mCherry-Gma</td>
			<td colspan="2">actin binding</td>
			<td colspan="2">UAS line</td>
			<td colspan="2">-</td>
			<td colspan="2">Gma fused to mCherry</td>
			<td>17</td>
		</tr>
		<tr>
			<td colspan="2">UAS-Lifeact-GFP</td>
			<td colspan="2">actin binding</td>
			<td colspan="2">UAS line</td>
			<td colspan="2">BL 35544</td>
			<td colspan="2"></td>
			<td>18</td>
		</tr>
		<tr>
			<td colspan="2">UAS-Lifeact-Ruby</td>
			<td colspan="2">actin binding</td>
			<td colspan="2">UAS line</td>
			<td colspan="2">BL 35545</td>
			<td colspan="2"></td>
			<td>18</td>
		</tr>
		<tr>
			<td colspan="2">UAS-CD8-GFP</td>
			<td colspan="2">membrane binding</td>
			<td colspan="2">UAS line</td>
			<td colspan="2">various stocks, e.g.: BL 32184</td>
			<td colspan="2"></td>
			<td>19</td>
		</tr>
		<tr>
			<td colspan="2">UAS-CD8-mCherry</td>
			<td colspan="2">membrane binding</td>
			<td colspan="2">UAS line</td>
			<td colspan="2">BL 27391 and 27392</td>
			<td colspan="2"></td>
			<td>20</td>
		</tr>
		<tr>
			<td colspan="2">UAS-palm-mCherry</td>
			<td colspan="2">membrane binding through palmitoylation</td>
			<td colspan="2">UAS line</td>
			<td colspan="2">BL 34514</td>
			<td colspan="2">UAS-brainbow</td>
			<td>21</td>
		</tr>
	</tbody>
</table>
Table 1: Fluorescently tagged protein markers suitable for studying muscle-tendon morphogenesis in vivo.
Here, we describe in detail how imaging of muscle-tendon morphogenesis in living pupae can be performed easily and successfully (Figure 1). Alternatively, pupae can be fixed, dissected and immunostained, which allows using antibodies to also label proteins for which no live markers are available22. In this case, the imaging quality is generally higher because there is no movement and the structure of interest can be placed in close proximity to the coverslip. However, dissection and fixation can lead to damage and molecular or tissue dynamics, for example, muscle twitching, can only be studied in the living organism.

<table><tbody><tr><td>Stereomicroscope</td><td>Leica</td><td>MZ6</td><td>product has been replaced by Leica M60</td></tr><tr><td>fly food in bottles (or vials)</td><td>-</td><td>-</td><td>standard culture medium</td></tr><tr><td>paint brush</td><td>da Vinci</td><td>1526Y</td><td>size 1</td></tr><tr><td>microscope slides</td><td>Thermo Scientific</td><td>VWR: 631-1303</td><td>76 x 26 mm</td></tr><tr><td>double-sided tape (optional)</td><td>Scotch</td><td>6651263</td><td>12 mm x 6.3 m</td></tr><tr><td>petri dishes</td><td>Greiner Bio-One</td><td>632102</td><td>94 x 16 mm</td></tr><tr><td>paper tissues</td><td>Th.Geyer</td><td>7695251</td><td></td></tr><tr><td>forceps #5 (Dumont, inox, standard)</td><td>Fine Science Tools</td><td>11251-20</td><td>0.1 mm x 0.06 mm tip</td></tr><tr><td>forceps #5 (Dumont, inox, biology grade)</td><td>Fine Science Tools</td><td>11252-20</td><td>0.05 mm x 0.02 mm tip</td></tr><tr><td>Cohan-Vannas spring scissors</td><td>Fine Science Tools</td><td>15000-02</td><td>straight tip</td></tr><tr><td>plastic slides with a groove (reusable)</td><td>custom-built</td><td>-</td><td>75 x 26 x 4 mm plexi glass slide with 1.0-1.5 mm wide and 0.3-0.4 mm deep groove</td></tr><tr><td>coverslips</td><td>Marienfeld</td><td>107032</td><td>18 x 18 mm, No. 1.5H</td></tr><tr><td>glycerol</td><td>Sigma-Aldrich</td><td>49781</td><td>dilute to 50 % in water</td></tr><tr><td>adhesive tape</td><td>Tesa</td><td>57370-02</td><td>1.5 mm x 10 m</td></tr></tbody></table>

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.

Various tissues can be imaged in vivo in developing fly pupae, making them an ideal model system to study morphogenesis of adult organs. Among these are the indirect flight muscles, the thorax epithelium including the tendon cells, the wing epithelium, abdominal muscles and the heart22,23,24,25,26,27. Here, we focus on live imaging of muscle and tendon morphogenesis. For a detailed description of indirect flight muscle and abdominal muscle morphogenesis and additional methods for studying muscle biology in Drosophila, we refer the reader to Weitkunat and Schnorrer 201422.
Live imaging of indirect flight muscles
For studying the long-term development of the indirect flight muscles consisting of the dorsolongitudinal muscles (DLMs) and the dorsoventral muscles (DVMs), globular Moesin actin-binding domain tagged with GFP (UAS-GFP-Gma) was expressed in all muscles using the Myocyte enhancer factor 2 (Mef2)-GAL4 driver (Figure 2A-D, Movie S1). Prior to imaging, a window in the pupal case was opened above the thorax as shown in Figure 1F. On a two-photon microscope, z-stacks were acquired every 20 min for 21 h starting at ≈11 h after puparium formation (11 h APF). In this time frame, the DLMs (green overlays in Figure 2A'-D') first initiate attachment to the tendon cells (Figure 2A) and then split while the muscle attachments mature (Figure 2B). Next, the myotubes shorten (Figure 2C) until they finally reach the maximally compacted stage at 30 h APF (Figure 2D). Taken together, this movie highlights the dramatic changes in muscle morphology that occur on the time scale of hours.
<img alt="Time-lapse microscopy, Drosophila indirect flight muscle, GFP-tagged, developmental biology study." class="xfigimg" src="/files/ftp_upload/57312/57312fig2.jpg" data-altupdatedat="1762215414000" data-alt="Figure 2"> 
Figure 2: Live imaging of indirect flight muscle and tendon morphogenesis. (A-D) Time points from a two-photon movie (Movie S1) using Mef2-GAL4, UAS-GFP-Gma as a marker for actin in the indirect flight muscles consisting of the dorsolongitudinal muscles (DLMs, highlighted in green in A'-D') and the dorsoventral muscles (DVMs, highlighted in blue in A'-D'). Scale bars are 100 µm. (E-H) Time points from a two-photon movie (Movie S2) using Duf-GAL4, UAS-CD8-GFP as a marker for indirect flight muscles and the thorax epithelium including tendon cells. Panels E'-H' show an overlay with a model of the muscle-tendon system at each time point, highlighting the long cellular extensions formed by the tendon epithelium (magenta) in contact with the muscles (green). Scale bars are 100 µm (estimated from size of imaged structures). (I-L) Time points from a two-color, spinning disc confocal movie (Movie S3) focusing on muscle-tendon attachment initiation. The tendon cells are labeled with sr-GAL4, UAS-palm-mCherry (magenta) and the dorsolongitudinal indirect flight muscles with Mhc-Tau-GFP (green). Scale bars are 10 µm. Time is indicated as hh:mm after puparium formation (APF). Note that not all movies were acquired on a temperature-controlled stage, therefore, developmental timing may diverge. <a href="https://cloudflare.jove.com/files/ftp_upload/57312/57312fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
For studying tendon and muscle morphogenesis at the same time, membrane-bound GFP (UAS-CD8-GFP) was expressed using Dumbfounded (Duf)-GAL4 as a driver, which is expressed both in the tendon cells and the indirect flight muscles (Figure 2E-H, Movie S2). A z-stack was taken on a two-photon microscope every 20 min starting at 16 h APF for 20 h. While the myotubes compact (green overlay in Figure 2E'-H'), the tendon cells form long cellular extensions that elongate with time (magenta overlay in Figure 2E'-H'). Taken together, this movie highlights the close interplay between the tendon and muscle cells in vivo.
To investigate muscle-tendon interaction in more detail, two-color, high-magnification imaging was performed. Pupae with myosin heavy chain (Mhc)-Tau-GFP in the DLMs and UAS-palmitoylated-mCherry (UAS-palm-mCherry) driven by stripe (sr)-GAL4 in the tendons were imaged every 5 min starting at 12 h APF for 4.5 h on a spinning disc confocal microscope (Figure 2I-L, Movie S3). At 12 h APF, the myotubes migrate towards their tendon target cells while forming long filopodia at their tips (Figure 2I). Subsequently, the muscle and tendon tissues interdigitate (Figure 2J, K) to form a stable attachment. As the attachment matures, fewer filopodia form and the muscle-tendon interface smoothens (Figure 2L). Thus, two-color, high-magnification imaging can be used to reveal cellular dynamics in detail in the living organism.
Live imaging of abdominal muscles
For live imaging of the abdominal muscles (Figure 3, Movie S4), Mef2-GAL4&gt;UAS-CD8-GFP was used as a marker and a window was opened above the abdomen in the pupal case as detailed in Figure 1G. Similar to the indirect flight muscle movie represented in Figure 2A-D, the formation and growth of the abdominal muscles can be followed over many hours of development (55 h in Movie S4). During this time, the myoblasts fuse to form growing myotubes (Figure 3A). The myotube tips migrate to their tendon targets, and after attaching to the tendon cells, the contractile units of the muscles, the sarcomeres, are formed (Figure 3B-D).
<img alt="Time-lapse microscopy of abdominal muscles; Mef2-GAL4, UAS-CD8-GFP fluorescence expression." class="xfigimg" src="/files/ftp_upload/57312/57312fig3.jpg" data-altupdatedat="1762215414000" data-alt="Figure 3"> 
Figure 3: Live imaging of abdominal muscle morphogenesis. (A-D) Time points from a movie (Movie S4) using Mef2-GAL4, UAS-CD8-GFP as a marker to follow abdominal muscle morphogenesis. Panels A'-E' show overlays with models of an abdominal muscle set (green) that forms de novo during the pupal stage. Scale bars are 100 µm. Time is indicated as hh:mm APF. The movie was acquired at room temperature. <a href="https://cloudflare.jove.com/files/ftp_upload/57312/57312fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Live imaging of twitching muscles
In contrast to Figure 2 and Figure 3, Figure 4 shows muscle dynamics occurring on the time scale of seconds: the live recording of muscle contractions. Pupae expressing βPS-Integrin-GFP under endogenous control were imaged with a time resolution of 0.65 s in a single z-plane on a confocal microscope (Movie S5). In the example displayed in Figure 4, the attachment sites of three DVMs (Figure 4A) were imaged for 10 min starting at 42 h APF. During this time, five twitch events were observed (Figure 4B-F), showing that the sarcomeres are already assembled well enough to support coordinated contractions at this time point in development. Hence, imaging of muscle twitching can be used as a functional read-out for sarcomerogenesis already during indirect flight muscle development28, as opposed to for example flight tests, which can only be performed after eclosion.
<img alt="Fluorescence microscopy of βPS-Integrin-GFP in dorsoventral muscles showing time-lapse attachment changes." class="xfigimg" src="/files/ftp_upload/57312/57312fig4.jpg" data-altupdatedat="1762215414000" data-alt="Figure 4"> 
Figure 4: Live imaging of twitching muscles. (A) First time-point from a movie (Movie S5) showing twitching of dorsoventral indirect flight muscles at 42 h APF using βPS-Integrin-GFP as a marker. In the field of view are the muscle attachment sites of DVM II 2, DVM III 1 and DVM III 2. (B-F) Color overlays of five individual twitch events, each showing the frame before the twitch (magenta) and the first frame of the twitch event (green). Note that the individual muscle fibers twitch independently from each other. Arrows highlight the twitching motion. The time resolution of the movie is 0.65 s. Scale bars are 25 µm. <a href="https://cloudflare.jove.com/files/ftp_upload/57312/57312fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Live imaging of endogenously tagged proteins
Similar to βPS-Integrin-GFP, a large collection of fusion proteins expressed in Drosophila under endogenous control has been generated2,3,4. These fly lines can be used to study for example the subcellular localization or the expression profiles of proteins of interest. The collection is especially helpful if specific antibodies are not available or protein dynamics need to be investigated in vivo without fixation. Figure 5 shows three examples of endogenously expressed fusion proteins from the fly TransgeneOme (fTRG) library, Hu li tai shao (Hts)-GFP (Figure 5A, B), Talin-GFP (Figure 5C, D) and βTubulin60D-GFP (Figure 5E, F). The expression of these proteins can be studied in all tissues that express them naturally. Here, we show the thorax epithelium (Figure 5A, C, E) and the indirect flight muscles or their attachment sites (Figure 5B, D, F) as examples.
<img alt="fluorescence microscopy images, GFP-tagged proteins in Drosophila epithelium, muscle attachment study" class="xfigimg" src="/files/ftp_upload/57312/57312fig5.jpg" data-altupdatedat="1762215414000" data-alt="Figure 5"> 
Figure 5: Live imaging of endogenously tagged proteins. (A, B) Maximum projections of z-stacks taken of pupae expressing Hts-GFP. (C, D) Maximum projections of z-stacks taken of pupae expressing Talin-GFP. (E, F) Maximum projections of z-stacks taken of pupae expressing βTubulin60D-GFP. Panels A, C and E show the thorax epithelium at 18, 24 and 18 h APF, respectively, and panels B, D and F show the indirect flight muscles or their attachment sites at 30 h APF. Scale bars are 25 µm. <a href="https://cloudflare.jove.com/files/ftp_upload/57312/57312fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae at JoVE.com

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.

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 ...

The overall goal of this procedure is to study muscle-tendon morphogenesis in living Drosophila pupae.This method can help answer key questions in the muscle development field.Such as how muscle cells interact with their tendon targets, and when they acquire the ability to twitch.The main advantage of this method is that muscle-tendon morphogenesis can be studied in the intact organism, and over many hours.To begin the protocol, use a wet brush to collect white pre-pupae from the walls of the bottles of fly stocks, and transfer them to a glass slide.Next, under a stereo microscope, remove clumps of fly food sticking to the pupae with a wet brush, and orient the ventral side of the pupae towards the glass slide.Discard the pupae that have started to turn brown, or that are still moving.Label the glass slide with the genotype, date and time of collection.Transfer the slide to a Petri dish and add a wet tissue to prevent the pupae from drying out.Store the pupae in an incubator at 25

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Research

JoVE Journal

Developmental Biology

11.7K Views.  Max Planck Institute of Biochemistry. The overall goal of this procedure is to study muscle-tendon morphogenesis in living Drosophila pupae.This method can help answer key questions in the muscle development field.Such as how muscle cells interact with their tendon targets, and when they acquire the ability to twitch.The main advantage of this method is that muscle-tendon morphogenesis can be studied in the intact organism, and over many hours.To begin the protocol, use a wet brush to collect white pre-pupae from the walls of the...

Video: In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Biology

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present methods for the development of assays to query potentially clinically significant nonsynonymous changes using in vivo complementation in zebrafish. Zebrafish (Danio rerio) are a useful animal system due to their experimental tractability; embryos are transparent to enable facile viewing, undergo rapid development ex vivo, and can be genetically manipulated.1 These aspects have allowed for significant advances in the analysis of embryogenesis, molecular processes, and morphogenetic signaling. Taken together, the advantages of this vertebrate model make zebrafish highly amenable to modeling the developmental defects in pediatric disease, and in some cases, adult-onset disorders. Because the zebrafish genome is highly conserved with that of humans (~70% orthologous), it is possible to recapitulate human disease states in zebrafish. This is accomplished either through the injection of mutant human mRNA to induce dominant negative or gain of function alleles, or utilization of morpholino (MO) antisense oligonucleotides to suppress genes to mimic loss of function variants. Through complementation of MO-induced phenotypes with capped human mRNA, our approach enables the interpretation of the deleterious effect of mutations on human protein sequence based on the ability of mutant mRNA to rescue a measurable, physiologically relevant phenotype. Modeling of the human disease alleles occurs through microinjection of zebrafish embryos with MO and/or human mRNA at the 1-4 cell stage, and phenotyping up to seven days post fertilization (dpf). This general strategy can be extended to a wide range of disease phenotypes, as demonstrated in the following protocol. We present our established models for morphogenetic signaling, craniofacial, cardiac, vascular integrity, renal function, and skeletal muscle disorder phenotypes, as well as others.

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Here, we present a systematic approach for developing physiologically relevant, sensitive and specific in vivo assays for interpreting variation in human pathology. Transient genetic manipulation via microinjection of WT and mutant human mRNA and morpholino (MO) antisense oligonucleotides harness the tractability of the developing zebrafish embryo to rapidly assay pathogenic mutations, especially, but not exclusively, in the context of human developmental disorders.

Here,  we present methods for the development of assays to query potentially clinically  significant nonsynonymous changes using in  vivo complementation ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

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 ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

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

Detection and Quantification of Single mRNA Molecules in Drosophila

Studying Muscle Transcriptional Dynamics at Single-molecule Scales in Drosophila

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

Author Spotlight: Investigating mRNA Spatial Distribution in Drosophila Muscle Tissue 

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

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

Viewing Microtubule Networks in Drosophila Neuromuscular Junctions and Muscle Cells

Using Drosophila Larval Neuromuscular Junction and Muscle Cells to Visualize Microtubule Network

The microtubule network is an essential component of the nervous system. Mutations in many microtubules regulatory proteins are associated with neurodevelopmental disorders and neurological diseases, such as microtubule-associated protein Tau to neurodegenerative diseases, microtubule severing protein Spastin and Katanin 60 cause hereditary spastic paraplegia and neurodevelopmental abnormalities, respectively. Detection of microtubule networks in neurons is advantageous for elucidating the pathogenesis of neurological disorders. However, the small size of neurons and the dense arrangement of axonal microtubule bundles make visualizing the microtubule networks challenging. In this study, we describe a method for dissection of the larval neuromuscular junction and muscle cells, as well as immunostaining of &#945;-tubulin and microtubule-associated protein Futsch to visualize microtubule networks in Drosophila melanogaster. The neuromuscular junction permits us to observe both pre-and post-synaptic microtubules, and the large size of muscle cells in Drosophila larva allows for clear visualization of the microtubule network. Here, by mutating and overexpressing Katanin 60 in Drosophila melanogaster,&#160;and then examining the microtubule networks in the neuromuscular junction and muscle cells, we&#160;accurately reveal the regulatory role of Katanin 60 in neurodevelopment. Therefore, combined with the powerful genetic tools of Drosophila&#160;melanogaster, this protocol greatly facilitates genetic screening and microtubule dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

Author Spotlight: Understanding Microtubule Network in Drosophila Neuromuscular Junctions 

Here, we present a detailed protocol to visualize the microtubule networks in neuromuscular junctions and muscle cells. Combined with the powerful genetic tools of Drosophila melanogaster, this protocol greatly facilitates genetic screening and microtubule-dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

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Explore More Videos

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

In Vivo Modeling of the Morbid Human Genome using Danio rerio

Imaging Through the Pupal Case of Drosophila melanogaster

Live-imaging of the Drosophila Pupal Eye

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

Studying Muscle Transcriptional Dynamics at Single-molecule Scales in Drosophila

Using Drosophila Larval Neuromuscular Junction and Muscle Cells to Visualize Microtubule Network