The authors would like to thank members of the Translational Genomics Group and Kathryn J Hanson for the feed-back and the improvements on this protocol. This project was carried out with research grant SAF2015-64500-R, which includes European Regional Development Funds, awarded to R.A by the Ministerio de Economia y Competitividad.

1. Fixation and resin embedding
<ol>
	<li>Anesthetize the flies with carbon dioxide (CO2) or via hypothermia using an ice block. Use a dissecting microscope (with low magnification to see the whole fly) and scissors to remove the legs, wings, head and the terminal part of abdomen to facilitate the penetration of the fixative. Careful handling of the carcasses is required at this step to avoid the deformation of the thorax. 
	NOTE: Start with at least 12 flies per genotype to ensure a sufficient number of properly processed individuals at the end of the procedure.</li>
	<li>Transfer the carcasses to a 1.5 mL tube on ice (placing a maximum of 6 individuals per tube) containing 200 &#181;L of solution 1. 
	NOTE: The carcasses cannot remain in the solution 1 for more than 20 min.
	<ol>
		<li>For solution 1, mix components in these volume proportions: &#188; 4% paraformaldehyde, &#188; 8% glutaraldehyde, &#188; 0.2 M Na2HPO4 and &#188; 0.2 M NaH2PO4.</li>
	</ol>
	</li>
	<li>Add 200 &#181;L of solution 2 and incubate on ice for 30 min.
	<ol>
		<li>Solution 2: Mix solution 1 and OsO4 in a 1:1 (v/v) proportion. 
		CAUTION: OsO4 is extremely toxic. Manipulate it in a fume hood with appropriate safety and disposal measures.</li>
	</ol>
	</li>
	<li>Remove all the liquid. Add 200 &#181;L of solution 2 and incubate on ice for 1-2 h.</li>
	<li>Remove the solution and dehydrate the samples through a graded ethanol series as follows: once in 30 %, 50%, 70%, 90%, and twice in absolute ethanol; 5 min each. Add 1 mL of ethanol to cover the samples. 
	NOTE: Once the tissue is placed in 90% ethanol, the subsequent steps can be performed at room temperature.</li>
	<li>Incubate the samples twice in propylene oxide for 10 min each incubation. 
	CAUTION: Propylene oxide is toxic, use it in a fume hood.</li>
	<li>Replace the propylene oxide by a 1:1 (v/v) mixture of epoxy resin and propylene oxide and incubate at room temperature overnight. 
	CAUTION: The resin is extremely toxic in a liquid state.The resin is available as a set of four components (A, B, C and D).
	<ol>
		<li>For epoxy resin, add in a disposable beaker 54 g of resin (A), 44.5 g of hardener (B), 2.5 g of accelerator (C) and 10 g of plasticizer (D). Combine all the resin components in a fume hood. Store the resin in aliquots at -20 &#176;C (for up to 6 months) or at -80 &#176;C (for several years).</li>
		<li>To discard the resin, bake it at 70 &#176;C for 24 h because solid resin is not toxic. Discard all the previous solutions in the Heavy Metals Group according to appropriate international procedures, because solutions can contain traces of OsO4.</li>
	</ol>
	</li>
	<li>Replace the previous mixture with 100% resin, and incubate the samples for 4 h.</li>
	<li>Transfer the carcasses to plastic molds containing new resin (one carcass in each well of the mold). Use a sharpened wooden stick or needle to transfer and orient the carcasses in the wells and leave the resin to polymerize overnight in a Pasteur oven at 70 &#176;C. 
	NOTE: To ensure appropriate orientation of the carcasses for posterior trimming and sectioning, the carcasses should be lying on their long side, with the anterior part of the carcass very near to the edge of the well.</li>
</ol>
2. Trimming and Sectioning of Blocks
<ol>
	<li>Remove the polymerized resin blocks from the molds. Use razor blades to trim away the resin, form a trapezoidal shape surrounding the carcass and give appropriate orientation in the microtome. Start sectioning all the carcasses after the transverse suture to obtain comparable results. 
	NOTE: The section must be perpendicular to the anteroposterior axis of the DLMs to trim the sample correctly ( Figure 3).</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56179/56179fig3.jpg" /> 
Figure 3. Dorsoventral sections of resin-embedded adult thoraces of the Myotonic Dystrophy model flies. In (A), the trimming of the sample was not correct as the section in the dorsal-right part of the image was not completely transversal to the muscles. As a consequence, the three muscle bundles on the upper right side seem bigger than the ones of the left side. This mistake would result in an overestimation of the muscle area. (B)&#160;Section from a correctly trimmed resin block. Scale bar: 200 &#181;m.
<ol>
	<li>Cut 1.5-&#181;m thick sections with an ultramicrotome and transfer the sections into water drops over gelatinized slides.
	<ol>
		<li>Obtain the sections from the mesothorax segment for all the samples to obtain comparable results.</li>
	</ol>
	</li>
	<li>Place the slides containing the sections on a heating block at 70-80 &#176;C until the H2O evaporates. At this point, the samples can be observed with the OsO4 contrast (Figure 4A).</li>
</ol>
3. Staining IFM sections
<ol>
	<li>Cover the sections with enough toluidine blue for approximately 30 s on the heating block.
	<ol>
		<li>For toluidine blue solution, dissolve 1% of toluidine blue and 1% of borax in H2O.</li>
	</ol>
	</li>
	<li>Rinse with H2O and leave the slides on a warm plate to evaporate the water. Repeat the staining with toluidine blue if the contrast needs to be enhanced (Figure 4B).</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56179/56179fig4.jpg" /> 
Figure 4. Dorsoventral sections of resin-embedded adult thoraces with different contrast stainings. Both panels show images of DM1 model flies fed with Abp1 as in Figure 2D. (A)&#160;Image of a section with the OsO4 contrast. (B)&#160;Section stained with toluidine blue allows better visualization of the muscle bundles. Scale bar: 200 &#181;m.
4. Mounting IFM sections
<ol>
	<li>Dry the slides with the heating block until the H2O evaporates.</li>
	<li>Put 1 or 2 drops of mounting medium on the sections and put on a coverslip. 
	Caution: Carry out this step in a fume hood because the mounting medium is toxic.</li>
</ol>
5. Acquisition of images and quantification of muscle area
<ol>
	<li>Take the images at low magnification so that the whole set of DLM muscles is in focus. We normally use 10X.
	<ol>
		<li>For statistical analysis take at least 5 serial images per fly and analyze at least 5 flies per experimental group.</li>
	</ol>
	</li>
	<li>Select the image containing the whole muscles (Figure 5A). Check the axis or orientation of the section and discard the samples with inappropriate orientation, which would distort the results (compare Figures 3A/ 3B to distinguish between badly and well-oriented sections, respectively).</li>
	<li>Use ImageJ software to select an area (in pixels) containing only the DLMs (Figure 5A/5B). For comparison between different experimental groups, the reference area to be used would always be the fly with the biggest muscles. In all the images of the different genotypes to compare, select an area containing DLMs of equal dimensions to the reference area (Figure 5B).</li>
	<li>Binarize the images with the Image-J command: Process/Binary/Make binary and delete the areas or pixels that do not correspond to the muscles of interest (Figure 5C).
	<ol>
		<li>In the case that artefactual black spots appear in areas other than muscle, delete them with the command Drawing tools and select the eraser symbol.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56179/56179fig5.jpg" /> 
Figure 5. Image analysis procedure for muscle area determination. (A)&#160;Transversal section of Myotonic Dystrophy type 1 model fly thorax with a rectangle containing the DLM. (B) Selected area with only the indirect flight muscles and the intestine (*). (C)&#160;Binarized image. The black areas correspond to the muscles of interest. The intestine has been erased for an accurate quantification of muscle area. Scale bar: 200 &#181;m.
<ol start="5">
	<li>Quantify percentage of pixels corresponding to muscle tissue (black pixels) out of total with the command: Analyze/Measurement and select the option area fraction. This percentage of area is an estimation of the DLM muscular mass of each fly.</li>
</ol>

<ol>
	<li>Fernandes, J., Bate, M., Vijayraghavan, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+indirect+flight+muscles+of+Drosophila.">Development of the indirect flight muscles of Drosophila.</a> Development. 113, 67-77 (1991).</li><li>Fernandes, J. J., Keshishian, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterning+the+dorsal+longitudinal+flight+muscles+(DLM)+of+Drosophila:+insights+from+the+ablation+of+larval+scaffolds.">Patterning the dorsal longitudinal flight muscles (DLM) of Drosophila: insights from the ablation of larval scaffolds.</a> Development. 122, 3755-3763 (1996).</li><li>Hartenstein, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atlas+of+Drosophila+Development.">Atlas of Drosophila Development.</a> Atlas Drosoph. Dev. , 1-57 (1993).</li><li>Tomlinson, A., Ready, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+fate+in+the+Drosophila+ommatidium.">Cell fate in the Drosophila ommatidium.</a> Dev. Biol. 123, 264-275 (1987).</li><li>Griffith, W. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Osmium+Tetroxide+And+Its+Applications.">Osmium Tetroxide And Its Applications.</a> Platin Met Rev. 18, 94-96 (1974).</li><li>Bird, T. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myotonic+Dystrophy+Type+1.">Myotonic Dystrophy Type 1.</a> GeneReviews. 1, 1-23 (1993).</li><li>Bargiela, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+autophagy+and+apoptosis+contribute+to+muscle+atrophy+in+a+myotonic+dystrophy+type+1+Drosophila+model.">Increased autophagy and apoptosis contribute to muscle atrophy in a myotonic dystrophy type 1 Drosophila model.</a> Dis Model Mech. 8, 679-690 (2015).</li><li>Llamusi, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscleblind,+BSF+and+TBPH+are+mislocalized+in+the+muscle+sarcomere+of+a+Drosophila+myotonic+dystrophy+model.">Muscleblind, BSF and TBPH are mislocalized in the muscle sarcomere of a Drosophila myotonic dystrophy model.</a> Dis. Model. Mech. 6, 184-196 (2012).</li><li>Garc&#237;a-L&#243;pez, A., Llamus&#237;, B., Orz&#225;ez, M., P&#233;rez-Pay&#225;, E., Artero, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+discovery+of+a+peptide+that+prevents+CUG-RNA+hairpin+formation+and+reverses+RNA+toxicity+in+myotonic+dystrophy+models.">In vivo discovery of a peptide that prevents CUG-RNA hairpin formation and reverses RNA toxicity in myotonic dystrophy models.</a> Proc. Natl. Acad. Sci. U.S.A. 108, 11866-11871 (2011).</li><li>van der Plas, M. C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Dystrophin+is+required+for+integrity+of+the+musculature.">Drosophila Dystrophin is required for integrity of the musculature.</a> Mech. Dev. 124, 617-630 (2007).</li><li>Lloyd, T. E., Taylor, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flightless+flies:+Drosophila+models+of+neuromuscular+disease.">Flightless flies: Drosophila models of neuromuscular disease.</a> Ann New York Acad Sci. , 1184 (2010).</li><li>Babcock, D. T., Ganetzky, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+improved+method+for+accurate+and+rapid+measurement+of+flight+performance+in+Drosophila.">An improved method for accurate and rapid measurement of flight performance in Drosophila.</a> J. Vis. Exp. , e51223 (2014).</li></ol>

No conflicts of interest declared.

It has been demonstrated that Drosophila melanogaster is a useful model to study human neuromuscular diseases7,10,11, including Myotonic Dystrophies, which are characterized by the appearance of muscular atrophy. The protocol presented here is a useful tool for quantifying the muscle degeneration caused by the onset or progression of a particular disease in a fly model. For example, it is also possible to follow and qua...

The thorax of the fruit fly contains two different classes of flight muscles, which are functionally, physiologically and anatomically distinct. These muscles are: the indirect flight muscles (IFM), which are composed of dorso-longitudinal (DLM) and dorso-ventral (DVM) muscles (Figure 1), and the synchronous flight control muscles1,2. These muscles together generate the elevated mechanical power required for flight. The size, distribution and rostro-caudal disposition of the IFMs allow an easy orientation for transversal sectioning3 (Figure 2A). For this reason, we have selected these muscles to study muscle atrophy in Drosophila melanogaster.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56179/56179fig1.jpg" /> 
Figure 1. Diagram of the thorax of the fruit fly showing the indirect flight muscles (IFMs) arrangement. (Left) represents a lateral view and (Right) represents a cross section of the thorax. The IFMs are composed of the Dorso-longitudinal (DLM) muscles (in red) and the Dorso-ventral (DVM) muscles (in green).
Preservation of tissue structure and the control over dorso-ventral axis orientation of histological sections are critical to ensure proper assessment of muscle cross-sectional area. To preserve muscle structure we used a fixation mixture modified from Tomlinson et al.4 . Moreover, because muscles are internal tissues, the impermeability of Drosophila&#39;s exoskeleton is a problem as fixation mixtures cannot penetrate to the target tissues. To circumvent this problem, we removed the fly head, legs, wings and the last two segments of the abdomen to create holes that allowed the fixation mixture to enter. As part of the fixation protocol we included treatment with osmium tetroxide (OsO4)5, which is extensively used because of its ability to fix fats, including triglycerides. OsO4 preserves most structures extremely well, particularly at the cytological level and at the same time provides contrast to the image. After fixation, Drosophila thoraces were embedded in resin for transversal semi-thin sectioning (1.5 &#181;m). For improved contrast, tissue can be additionally stained with toluidine blue. Images of complete thoraces were taken at 10X and muscle area was quantified by binarizing images (of equal dimensions) and quantifying percentage of pixels corresponding to muscle tissue (black pixels) out of total, with ImageJ software.
Modifications on the tissue preparation and fixation mixtures, as the increase of the concentration of OsO4 and glutaraldehyde solution, introduced in this protocol, allowed unique preservation of muscle tissue. This is because the protocol avoids the degradation and deformation of the tissue, making the posterior analysis of the samples more reliable&#160;even in highly atrophic conditions associated with neuromuscular degenerative diseases such as Myotonic Dystrophy (DM). In its more common form, DM type 1, this rare genetic disorder is brought about by expanded CUG repeats in myotonic dystrophy protein kinase (DMPK) transcripts. Mutant DMPK RNA aggregates form&#160;ribonuclear foci that sequester the Muscleblind-like nuclear RNA-binding proteins (MBNL1-3; Muscleblind (Mbl) in Drosophila)6. We generated a Drosophila model of Myotonic Dystrophy by expressing 250 CTG repeats under the muscular myosin heavy chain promoter (Mhc-Gal4). Model flies were flightless with a typical &#39;up-held wings&#39; phenotype and had serious muscle atrophy in their IFMs (Figure 2B). Previous studies performed in our laboratory have shown that determination of the muscle area of IFMs is a reliable method to quantify the effects of different chemical or genetic modifiers of the muscle atrophy in these model flies7. As an example, overexpression of Mbl isoform C in flies expressing the 250 CTG repeats in the muscle, achieved a rescue of muscle area, as Mbl depletion by sequestration is the triggering factor in DM1 pathogenesis8 (Figure 2C). Muscle area was also rescued after feeding the DM model flies with Abp1, a hexapeptide with proven anti-DM1 activity9 (Figure 2D).
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56179/56179fig2.jpg" /> 
Figure 2. Quantification of dorsoventral sections of resin-embedded adult thoraces. (A-D) Indirect flight muscles of Drosophila melanogaster with the indicated relevant genotypes. (A) Control flies (yw). (B) Expression of 250 non-coding CTG repeats in muscle (UAS-CTG(250)x) caused a reduction of muscle area in DLMs in comparison to control flies. (C) This muscle atrophy phenotype was rescued by overexpression of Muscleblind (MblC) (UAS-CTG(250)x UAS-MblC) and (D) feeding the model flies with the hexapeptide Abp1 (UAS-CTG(250)x Abp1). In all images the dorsal side is on top. Transgenes were driven to muscle using a Myosin heavy-chain promoter (Mhc)-Gal4. (E) Quantification of percentage of muscle area relative to the control flies&#160;confirmed that the differences were significant. The histogram shows means &#177; S.E.M. **p&#60;0.01 and *p&#60;0.05 (Student&#180;s t-test). Scale bar: 200 &#181;m. <a href="//cloudflare.jove.com/files/ftp_upload/56179/56179fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The method here reported will be of interest to researchers focusing on muscle development, maintenance and aging, disease pathology and drug testing as it provides reliable information about how muscle tissue responds to both endogenous and external factors.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Image-J software</td>
			<td>National Institutes of Health</td>
			<td></td>
			<td>https://imagej.nih.gov/ij/</td>
		</tr>
		<tr>
			<td>Ultramicrotome</td>
			<td>Leica</td>
			<td>Leica UC6</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Leica</td>
			<td>Leica MZ6</td>
			<td>Bright field technique.</td>
		</tr>
		<tr>
			<td>Razor blades</td>
			<td>Electron Microscopy Sciences</td>
			<td>71970</td>
			<td>Several alternative providers exist.</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>World Precision World</td>
			<td>14003</td>
			<td>Several alternative providers exist.</td>
		</tr>
		<tr>
			<td>Embedding molds</td>
			<td>Electron Microscopy Sciences</td>
			<td>70900</td>
			<td>Several alternative providers exist.</td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Fluka (Sigma)</td>
			<td>49624</td>
			<td>Toxic.</td>
		</tr>
		<tr>
			<td>OsO4</td>
			<td>Polyscience</td>
			<td>0972A</td>
			<td>Extremely toxic.</td>
		</tr>
		<tr>
			<td>Propylene oxide</td>
			<td>Sigma Aldrich</td>
			<td>82320-250ML</td>
			<td>Extremely toxic.</td>
		</tr>
		<tr>
			<td>resin (Durcupan)</td>
			<td>Sigma Aldrich</td>
			<td>44611-44614</td>
			<td>Carcinogenic when it is unpolymerized.</td>
		</tr>
		<tr>
			<td>Toluidine blue</td>
			<td>Panreac</td>
			<td>251176</td>
			<td>Toxic.</td>
		</tr>
		<tr>
			<td>Mountant Medium (DPX)</td>
			<td>Sigma Aldrich</td>
			<td>44581</td>
			<td>Dangerous.</td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Sigma Aldrich</td>
			<td>P6148-500G</td>
			<td>Harmful.</td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>Panreac</td>
			<td>122507</td>
			<td>0.2 M dilution.</td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Panreac</td>
			<td>121677</td>
			<td>0.2 M dilution.</td>
		</tr>
		<tr>
			<td>Borax</td>
			<td>Panreac</td>
			<td>3052</td>
			<td>Toxic.</td>
		</tr>
	</tbody>
</table>

optical cross-sectional muscle area determination of drosophila melanogaster adult indirect flight muscles

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

To quantify whether the overexpression of MblC or the administration of Abp1 had any effect in the atrophic phenotype of the Myotonic Dystrophy fly model we focused on the DLMs, which are part of IFMs (Figure 1). We determined that the model flies, which express 250 non-coding CTG repeats throughout the musculature driven by the Myosin heavy-chain promoter (Mhc)-Gal4, had a 50% reduction of muscle area compared to control flies. In contrast, co-expre...

Watch this Scientific Journal Video about Optical Cross-Sectional Muscle Area Determination of Drosophila Melanogaster Adult Indirect Flight Muscles at JoVE.com

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

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

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

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

optical-cross-sectional-muscle-area-determination-drosophila

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8.9K Views.  University of Valencia. We report a method to quantify muscle area, which is an indirect method to determine muscle mass in Drosophila adults. We demonstrate the application of our methodology by analyzing the indirect flight muscles in a Drosophila model of Myotonic Dystrophy disease.

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

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Visually Mediated Odor Tracking During Flight in Drosophila

Flying insects use visual cues to stabilize their heading in a wind stream. Many animals additionally track odors carried in the wind. As such, visual stabilization of upwind tracking directly aids in odor tracking. But do olfactory signals directly influence visual tracking behavior independently from wind cues? Additionally, recent advances in olfactory molecular genetics and neurophysiology have motivated novel quantitative behavioral analyses to assess the behavioral influence of (e.g.) genetically inactivating specific olfactory activation circuits. We modified a magnetic tether system originally devised for vision experiments by equipping the arena with narrow laminar flow odor plumes. Here we focus on experiments that can be performed after a fly is tethered and is able to navigate in the magnetic arena. We show how to acquire video images optimized for measuring body angle, how to judge stable odor tracking, and we illustrate two experiments to examine the influence of visual cues on odor tracking.

Here we describe how to optimize the acquired video image for an olfactory magnetic-tether (OMT) apparatus. We also describe two sample experimental protocols for studying visuo-olfactory fusion.

Flying insects use visual cues to stabilize their heading in a wind stream. Many animals additionally track odors carried in the wind. As such, visual ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

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

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

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

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

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function to maintain glucose homeostasis during developmental larval stages 1,2. Studies on largely post-mitotic adult flies have revealed perturbation of glucose homeostasis as the result of genetic ablation of insulin-like peptide (ILP) producing cells (IPCs) 3. Thus, adult fruit flies hold great promise as a suitable genetic model system for metabolic disorders including type II diabetes. To further develop the fruit fly system, comparable physiological assays used to measure glucose tolerance and insulin sensitivity in mammals must be established. To this end, we have recently described a novel procedure for measuring oral glucose tolerance response in the adult fly and demonstrated the importance of adult IPCs in maintaining glucose homeostasis 4,5. Here, we have modified a previously described procedure for insulin injection 6 and combined it with a novel hemolymph extraction method to measure peripheral insulin sensitivity in the adult fly. Uniquely, our protocol allows direct physiological measurements of the adult fly's ability to dispose of a peripheral glucose load upon insulin injection, a methodology that makes it feasible to characterize insulin signaling mutants and potential interventions affecting glucose tolerance and insulin sensitivity in the adult fly.

Insulin Injection and Hemolymph Extraction to Measure Insulin Sensitivity in Adult Drosophila melanogaster

Conserved insulin signaling pathways found in the fruit fly Drosophila melanogaster make this organism a potential tool for modeling metabolic disorders including type II diabetes. To this end, it is critical to establish physiological assays to effectively measure systemic insulin action in peripheral glucose disposal in the adult fly.

Conserved nutrient sensing mechanisms exist between mammal and fruit fly where peptides resembling mammalian insulin and glucagon, respectively function ...

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+ handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+ signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e., mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

We describe a method to directly measure muscle force, muscle power, contractile kinetics and fatigability of isolated skeletal muscles in an in vitro system using field stimulation. Valuable information on Ca2+ handling properties and contractile machinery of the muscle can be obtained using different stimulating protocols.

 
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Maintenance of a Drosophila melanogaster Population Cage

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life cycle of Drosophila enables collection of large quantities of material from embryos, larvae, pupae and adult flies, in a synchronized way, at a low economic cost. A major strategy for propagating large numbers of flies is the use of a fly population cage. This useful and common tool in the Drososphila community is an efficient way to regularly produce milligrams to tens of grams of embryos, depending on uniformity of developmental stage desired. While a population cage can be time consuming to set up, maintaining a cage over months takes much less time and enables rapid collection of biological material in a short period. This paper describes a detailed and flexible protocol for the maintenance of a Drosophila melanogaster population cage, starting with 1.5 g of harvested material from the previous cycle.

Maintenance of a Drosophila melanogaster Population Cage

This manuscript reports a detailed protocol for culturing, on a regular basis, a population of Drosophila melanogaster using a fly population cage.

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life ...