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 &#176;C (or 27 &#176;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 &#176;C or 27 &#176;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 &#176;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 &#181;L) of 50% glycerol solution above each pupal case opening onto the coverslip using a 20 &#181;L 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="Figure 1" class="xfigimg" src="/files/ftp_upload/57312/57312fig1.jpg" /> 
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 &#8776;11 h after puparium formation (11 h APF). In this time frame, the DLMs (green overlays in Figure 2A&#39;-D&#39;) 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="Figure 2" class="xfigimg" src="/files/ftp_upload/57312/57312fig2.jpg" /> 
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&#39;-D&#39;) and the dorsoventral muscles (DVMs, highlighted in blue in A&#39;-D&#39;). Scale bars are 100 &#181;m. (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&#39;-H&#39; 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 &#181;m (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 &#181;m. 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&#39;-H&#39;), the tendon cells form long cellular extensions that elongate with time (magenta overlay in Figure 2E&#39;-H&#39;). 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&#62;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="Figure 3" class="xfigimg" src="/files/ftp_upload/57312/57312fig3.jpg" /> 
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&#39;-E&#39; show overlays with models of an abdominal muscle set (green) that forms de novo during the pupal stage. Scale bars are 100 &#181;m. 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 &#946;PS-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="Figure 4" class="xfigimg" src="/files/ftp_upload/57312/57312fig4.jpg" /> 
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 &#946;PS-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 &#181;m. <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 &#946;PS-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 &#946;Tubulin60D-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="Figure 5" class="xfigimg" src="/files/ftp_upload/57312/57312fig5.jpg" /> 
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 &#946;Tubulin60D-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 &#181;m. <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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11.3K 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

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

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. ...

Using In Vivo and Tissue and Cell Explant Approaches to Study the Morphogenesis and Pathogenesis of the Embryonic and Perinatal Aorta

The aorta is the largest artery in the body. The aortic wall is composed of an inner layer of endothelial cells, a middle layer of alternating elastic lamellae and smooth muscle cells (SMCs), and an outer layer of fibroblasts and extracellular matrix. In contrast to the widespread study of pathological models (e.g., atherosclerosis) in the adult aorta, much less is known about the embryonic and perinatal aorta. Here, we focus on SMCs and provide protocols for the analysis of the morphogenesis and pathogenesis of embryonic and perinatal aortic SMCs in normal development and disease. Specifically, the four protocols included are: i) in vivo embryonic fate mapping and clonal analysis; ii) explant embryonic aorta culture; iii) SMC isolation from the perinatal aorta; and iv) subcutaneous osmotic mini-pump placement in pregnant (or non-pregnant) mice. Thus, these approaches facilitate the investigation of the origin(s), fate, and clonal architecture of SMCs in the aorta in vivo. They allow for modulating embryonic aorta morphogenesis in utero by continuous exposure to pharmacological agents. In addition, isolated aortic tissue explants or aortic SMCs can be used to gain insights into the role of specific gene targets during fundamental processes such as muscularization, proliferation, and migration. These hypothesis-generating experiments on isolated SMCs and the explanted aorta can then be assessed in the in vivo context through pharmacological and genetic approaches.

Using In Vivo and Tissue and Cell Explant Approaches to Study the Morphogenesis and Pathogenesis of the Embryonic and Perinatal Aorta

Protocols for studying the embryonic and perinatal murine aorta using in vivo clonal analysis and fate mapping, aortic explants, and isolated smooth muscle cells are detailed here. These diverse approaches facilitate the investigation of the morphogenesis of the embryonic and perinatal aorta in normal development and the pathogenesis in disease.

The aorta is the largest artery in the body. The aortic wall is composed of an inner layer of endothelial cells, a middle layer of alternating elastic ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analysis and Imaging

Multicellular tubes, fundamental units of all internal organs, are composed of polarized epithelial or endothelial cells, with apical membranes lining the lumen and basolateral membranes contacting each other and/or the extracellular matrix. How this distinctive membrane asymmetry is established and maintained during organ morphogenesis is still an unresolved question of cell biology. This protocol describes the C. elegans intestine as a&#160;model for the analysis of polarized membrane biogenesis during tube morphogenesis, with emphasis on apical membrane and lumen biogenesis. The C. elegans twenty-cell single-layered intestinal epithelium is arranged into a simple bilaterally symmetrical tube, permitting analysis on a single-cell level. Membrane polarization occurs concomitantly with polarized cell division and migration during early embryogenesis, but de novo polarized membrane biogenesis continues throughout larval growth, when cells no longer proliferate and move. The latter setting allows one to separate subcellular changes that simultaneously mediate these different polarizing processes, difficult to distinguish in most polarity models. Apical-, basolateral membrane-, junctional-, cytoskeletal- and endomembrane components can be labeled and tracked throughout development by GFP fusion proteins, or assessed by in situ antibody staining. Together with the organism&#39;s genetic versatility, the C. elegans intestine thus provides a unique in vivo model for the visual, developmental, and molecular genetic analysis of polarized membrane and tube biogenesis. The specific methods (all standard) described here include how to: label intestinal subcellular components by antibody staining; analyze genes involved in polarized membrane biogenesis by loss-of-function studies adapted to the typically essential tubulogenesis genes; assess polarity defects during different developmental stages; interpret phenotypes by epifluorescence, differential interference contrast (DIC) and confocal microscopy; quantify visual defects. This protocol can be adapted to analyze any of the often highly conserved molecules involved in epithelial polarity, membrane biogenesis, tube and lumen morphogenesis.

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analy

The transparent C. elegans intestine can serve as an "in vivo tissue chamber" for studying apicobasal membrane and lumen biogenesis at the single-cell and subcellular level during multicellular tubulogenesis. This protocol describes how to combine standard labeling, loss-of-function genetic/RNAi and microscopic approaches to dissect these processes on a molecular level.

Multicellular tubes, fundamental units of all internal organs, are composed of polarized epithelial or endothelial cells, with apical membranes lining the ...

The C. elegans Excretory Canal as a Model for Intracellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis in a Single Cell: labeling by GFP-fusions, RNAi Interaction Screen and Imaging

The four C. elegans excretory canals are narrow tubes extended through the length of the animal from a single cell, with almost equally far extended intracellular endotubes that build and stabilize the lumen with a membrane and submembraneous cytoskeleton of apical character. The excretory cell expands its length approximately 2,000 times to generate these canals, making this model unique for the in vivo assessment of de novo polarized membrane biogenesis, intracellular lumen morphogenesis and unicellular tubulogenesis. The protocol presented here shows how to combine standard labeling, gain- and loss-of-function genetic or RNA interference (RNAi)-, and microscopic approaches to use this model to visually dissect and functionally analyze these processes on a molecular level. As an example of a labeling approach, the protocol outlines the generation of transgenic animals with fluorescent fusion proteins for live analysis of tubulogenesis. As an example of a genetic approach, it highlights key points of a visual RNAi-based interaction screen designed to modify a gain-of-function cystic canal phenotype. The specific methods described are how to: label and visualize the canals by expressing fluorescent proteins; construct a targeted RNAi library and strategize RNAi screening for the molecular analysis of canal morphogenesis; visually assess modifications of canal phenotypes; score them by dissecting fluorescence microscopy; characterize subcellular canal components at higher resolution by confocal microscopy; and quantify visual parameters. The approach is useful for the investigator who is interested in taking advantage of the C. elegans excretory canal for identifying and characterizing genes involved in the phylogenetically conserved processes of intracellular lumen and unicellular tube morphogenesis.

The C. elegans Excretory Canal as a Model for Intracellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis in a Single Cell: labeling by GFP-fusions, RNAi Interaction Screen and Imaging

The C. elegans excretory canal is a unique single-cell model for the visual in vivo analysis of de novo polarized membrane biogenesis. This protocol describes a combination of standard genetic/RNAi and imaging approaches, adaptable for the identification and characterization of molecules directing unicellular tubulogenesis, and apical membrane and lumen biogenesis.

The four C. elegans excretory canals are narrow tubes extended through the length of the animal from a single cell, with almost equally far extended ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Cell Aggregation Assays to Evaluate the Binding of the Drosophila Notch with Trans-Ligands and its Inhibition by Cis-Ligands

Notch signaling is an evolutionarily conserved cell-cell communication system used broadly in animal development and adult maintenance. Interaction of the Notch receptor with ligands from neighboring cells induces activation of the signaling pathway (trans-activation), while interaction with ligands from the same cell inhibits signaling (cis-inhibition). Proper balance between trans-activation and cis-inhibition helps establish optimal levels of Notch signaling in some contexts during animal development. Because of the overlapping expression domains of Notch and its ligands in many cell types and the existence of feedback mechanisms, studying the effects of a given post-translational modification on trans- versus cis-interactions of Notch and its ligands in vivo is difficult. Here, we describe a protocol for using Drosophila S2 cells in cell-aggregation assays to assess the effects of knocking down a Notch pathway modifier on the binding of Notch to each ligand in trans and in cis. S2 cells stably or transiently transfected with a Notch-expressing vector are mixed with cells expressing each Notch ligand (S2-Delta or S2-Serrate). Trans-binding between the receptor and ligands results in the formation of heterotypic cell aggregates and is measured in terms of the number of aggregates per mL composed of &#62;6 cells. To examine the inhibitory effect of cis-ligands, S2 cells co-expressing Notch and each ligand are mixed with S2-Delta or S2-Serrate cells and the number of aggregates is quantified as described above. The relative decrease in the number of aggregates due to the presence of cis-ligands provides a measure of cis-ligand-mediated inhibition of trans-binding. These straightforward assays can provide semi-quantitative data on the effects of genetic or pharmacological manipulations on the binding of Notch to its ligands, and can help deciphering the molecular mechanisms underlying the in vivo effects of such manipulations on Notch signaling.

Cell Aggregation Assays to Evaluate the Binding of the Drosophila Notch with Trans-Ligands and its Inhibition by Cis-Ligands

Complexity of in vivo systems makes it difficult to distinguish between the activation and inhibition of Notch receptor by trans- and cis-ligands, respectively. Here, we present a protocol based on in vitro cell-aggregation assays for qualitative and semi-quantitative evaluation of the binding of Drosophila Notch to trans-ligands vs cis-ligands.

Notch signaling is an evolutionarily conserved cell-cell communication system used broadly in animal development and adult maintenance. Interaction of the ...

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 ...