We thank Gillian Stanfield for comments on the manuscript. T.A.J. is supported by the University of Utah Genetics Training Grant T32-GM007464 from NIH NIGMS.

1. Mosaic Generation
<ol>
	<li>Genetic Cross 
	w FRT19A tub:GAL80 FLP122/Y; Btl-GAL4, UAS-GFP (males) X * FRT19A/Bal (females), where * represents the mutant to be examined. Note: Set up cross at least one day prior to experiment, as this allows animals to mate appropriately.</li>
	<li>Pre-lay: Transfer cross to fly-food vials with a small smear of fresh yeast paste placed on the media. Allow to lay for 1 hr at 25 &deg;C.</li>
	<li>Lay: Transfer animals from pre-lay vial to a fresh vial with a small smear of fresh yeast paste. Allow to lay for 6 hr at 25 &deg;C.</li>
	<li>Transfer adults back to pre-lay vial for storage and use in subsequent experiments. Immediately heat shock the lay vial containing 0-6 hr old embryos for 45 min at 38 &deg;C, in a circulating water bath. Note: The surface of the food should be below the water level to verify all embryos are appropriately heat shocked.</li>
	<li>Incubate heat shocked vials at 25 &deg;C for 4-5 days until third instar larvae start to wander.</li>
</ol>
2. Screen for Mosaic Animals
<ol>
	<li>Using fine forceps gently pick wandering third-instar larvae from the sides of the vial and place them into chilled 100% glycerol in a small plastic plate.</li>
	<li>Examine larvae using a high magnification-dissecting microscope equipped with fluorescent optics. Identify larvae with mosaic expression of fluorescently labeled tracheal cells. Typically, this can be done at 10-20X magnification.</li>
</ol>
3. Heat Fixation
<ol>
	<li>Using fine forceps carefully pick mosaic animals out of the dish and place in a drop of fresh 100% glycerol on a 75x25x1 mm white glass microscope slide. Note: multiple animals can be placed in the drop (Figure 1A). 
	Animals can also be placed in a drop of glycerol directly onto a coverslip, but care must be taken during fixation (step 3.2), as the animals will heat very quickly and can become over-fixed.</li>
	<li>Place glass slide onto 70 &deg;C heat block until the animals just stop moving, no more than 20 sec for a slide, and 10 sec for a cover slip (Figure 1B). Animals will wriggle initially but stop and become rigid and elongated once dead. Note: overlong exposure to heat will damage terminal branches and lumens as well as make the GFP signal dim and diffuse. For optimal efficiency it is recommended to have the heat block in the same room as the microscopes.</li>
	<li>Using fine forceps carefully orient all larvae in the drop in a single direction, such that all larvae on the slide are oriented parallel to one another (Figure 1C).</li>
	<li>Gently place an 18x18mm micro cover glass on top of larvae (or invert the coverslip onto a glass slide) and avoid forming bubbles (Figure 1D). Note: coverslip may not lie exactly flat over larvae, this is not a problem.</li>
	<li>Image larvae within 30 min. Heat fixed terminal cells will degrade and GFP signal will become very diffuse.</li>
</ol>
4. In vivo Imaging of Tracheal Terminal Cells
<ol>
	<li>Place slide on the stage of a compound stereomicroscope. Using a 5X objective locate the two parallel dorsal trunks running from anterior to posterior on the dorsal surface of the animal (Figure 1F). Locate the two dorsal terminal cells, directly between the two dorsal trunks. This helps identify the appropriate segment for analysis.</li>
	<li>Once oriented, rotate the animals to image desired terminal cells. To do this, carefully push the coverslip on the edge with the forceps perpendicular to the long axis of the larvae, slowly rolling the larvae underneath the coverslip (Figures 1E and 1G).</li>
	<li>Selecting the appropriate terminal cells is critical for consistent results and comparisons between mutants. The fat body branch (FB) terminal cells are easily found adjacent to the dorsal trunk, lateral to the midline. The lateral group G terminal cells (LG) are easily found and are located just lateral of the ventral midline (Figure 1H). Note: in the most anterior (Tr1) and posterior (Tr10) segments the arrangement of terminal cell is divergent from other segments. We only analyze terminal cells in segments Tr2-Tr9. Dorsal terminal cells have a more stereotyped branching pattern than other tracheal terminal cells. This may depend on additional, non-cell autonomous signals, and this should be considered when deciding whether to include them in analysis.</li>
	<li>Once a terminal cell of interest has been identified, capture an image using the desired magnification; typically 10X or 20X is best (Figure 2A). Note: due to variable branching patterns, try to capture images that include as many branches and branch tips as possible.</li>
	<li>Without moving the stage, switch off the fluorescence and turn on the transmitted light to capture a brightfield image of the same cell and focal plane (Figure 2B). The lumen will be visualized as darkly contrasting within the terminal cell space. If planning on quantify terminal cell branches and lumens, collect multiple images in multiple focal planes of each terminal cell.</li>
</ol>
5. Analysis and Quantification of Terminal Cell Morphology Using ImageJ
<ol>
	<li>Open the fluorescent and brightfield images using the ImageJ (<a href="http://rsb.info.nih.gov/ij/" target="_blank">http://rsb.info.nih.gov/ij/</a>) plug-in NeuronJ (<a href="http://www.imagescience.org/meijering/software/neuronj/" target="_blank">http://www.imagescience.org/meijering/software/neuronj/</a>).</li>
	<li>Using the "Add tracings" tool, manually trace the branches and lumen of the entire terminal cell.</li>
	<li>Using the "Label tracings" tool, rename and recolor to assign each line within the trace a specific designation.</li>
	<li>Using the "Set parameters" tool, adjust the line width from 6-10 pixels (for 512x512 images obtained from the digital camera connected to the compound microscope. For higher or lower resolution cameras, the pixel width should be scaled appropriately), using all other default settings, and choose "OK". Then use the "Make snapshot" tool to take an image. Choose "Draw trace" to get a snapshot of the trace that can be shown side-by-side with the original.</li>
	<li>Select the "Measure tracings" tool and select the tracing type (i.e. which branches to measure based on the specific designations assigned in step 5.3). Next, select "Display tracing measurements" and "Clear previous measurements" then click "RUN". This will produce a spreadsheet that contains the name, label, length (in pixels), and other data for the image. This data can then be saved as a Microsoft Excel spreadsheet for further statistical analysis.</li>
</ol>

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	<li>Manning, G., Krasnow, M. A. <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+Drosophila+Tracheal+System.">Development of the Drosophila Tracheal System.</a> The Development of Drosophila melanogaster. , 609-685 (1993).</li><li>Jarecki, J., Johnson, E., Krasnow, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxygen+regulation+of+airway+branching+in+Drosophila+is+mediated+by+branchless+FGF.">Oxygen regulation of airway branching in Drosophila is mediated by branchless FGF.</a> Cell. 99 (2), 211-220 (1999).</li><li>Ghabrial, A. S., Krasnow, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Social+interactions+among+epithelial+cells+during+tracheal+branching+morphogenesis.">Social interactions among epithelial cells during tracheal branching morphogenesis.</a> Nature. 441 (7094), 746-749 (2006).</li><li>Ruiz, O. E., Nikolova, L. S., Metzstein, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Zpr1+(Zinc+Finger+Protein+1)+Is+Required+Downstream+of+Both+EGFR+And+FGFR+Signaling+in+Tracheal+Subcellular+Lumen+Formation.">Drosophila Zpr1 (Zinc Finger Protein 1) Is Required Downstream of Both EGFR And FGFR Signaling in Tracheal Subcellular Lumen Formation.</a> PLoS ONE. 7 (9), e45649 (2012).</li><li>Ghabrial, A. S., Levi, B. P., Krasnow, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+systematic+screen+for+tube+morphogenesis+and+branching+genes+in+the+Drosophila+tracheal+system.">A systematic screen for tube morphogenesis and branching genes in the Drosophila tracheal system.</a> PLoS Genetics. 7 (7), e1002087 (2011).</li><li>Baer, M. M., Bilstein, A., Leptin, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+clonal+genetic+screen+for+mutants+causing+defects+in+larval+tracheal+morphogenesis+in+Drosophila.">A clonal genetic screen for mutants causing defects in larval tracheal morphogenesis in Drosophila.</a> Genetics. 176 (4), 2279-2291 (2007).</li><li>Schottenfeld-Roames, J., Ghabrial, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whacked+and+Rab35+polarize+dynein-motor-complex-dependent+seamless+tube+growth.">Whacked and Rab35 polarize dynein-motor-complex-dependent seamless tube growth.</a> Nat. Cell Biol. 14 (4), 386-393 (2012).</li><li>Levi, B. P., Ghabrial, A. S., Krasnow, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+talin+and+integrin+genes+are+required+for+maintenance+of+tracheal+terminal+branches+and+luminal+organization.">Drosophila talin and integrin genes are required for maintenance of tracheal terminal branches and luminal organization.</a> Development. 133 (12), 2383-2393 (2006).</li><li>Jones, T. A., Metzstein, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+function+for+the+PAR+complex+in+subcellular+morphogenesis+of+tracheal+terminal+cells+in+Drosophila+melanogaster.">A novel function for the PAR complex in subcellular morphogenesis of tracheal terminal cells in Drosophila melanogaster.</a> Genetics. 189 (1), 153-164 (2011).</li><li>Gervais, L., Casanova, J. <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+coupling+of+cell+elongation+and+lumen+formation+in+a+single+cell.">In vivo coupling of cell elongation and lumen formation in a single cell.</a> Curr. Biol. 20 (4), 359-366 (2010).</li><li>Lee, T., Luo, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mosaic+analysis+with+a+repressible+cell+marker+for+studies+of+gene+function+in+neuronal+morphogenesis.">Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis.</a> Neuron. 22 (3), 451-461 (1999).</li><li>Shiga, Y., Tanaka-Matakatsu, M., Hayashi, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+nuclear+GFP/&#223;-galactosidase+fusion+protein+as+a+marker+for+morphogenesis+in+living+Drosophila.">A nuclear GFP/&#223;-galactosidase fusion protein as a marker for morphogenesis in living Drosophila.</a> Dev. Growth Differ. 38, 99-106 (1996).</li><li>Meijering, E., Jacob, M., Sarria, J. -. C. F., Steiner, P., Hirling, H., Unser, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+and+validation+of+a+tool+for+neurite+tracing+and+analysis+in+fluorescence+microscopy+images.">Design and validation of a tool for neurite tracing and analysis in fluorescence microscopy images.</a> Cytometry. Part A: The Journal of the International Society for Analytical Cytology. 58 (2), 167-176 (2004).</li><li>Centanin, L., Dekanty, A., Romero, N., Irisarri, M., Gorr, T. A., Wappner, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+autonomy+of+HIF+effects+in+Drosophila:+tracheal+cells+sense+hypoxia+and+induce+terminal+branch+sprouting.">Cell autonomy of HIF effects in Drosophila: tracheal cells sense hypoxia and induce terminal branch sprouting.</a> Dev. Cell. 14 (4), 547-558 (2008).</li></ol>

The authors declare that they have no competing financial interest.

The heat fixation technique described here is a rapid and convenient tool for imaging Drosophila larval tracheal terminal cells. Here, we use this technique to examine the branching and lumen pattern of wild-type cells. Tracheal cells expressing GFP, driven by the tracheal specific promoter breathless, can be easily visualized through the larval cuticle after heat fixation. The specific branching patterns of individual terminal cells, as well as those of the air-filled lumen, can be quickly visualized and measured using this method. This technique can also be performed on both first and second instar larvae. However, care must be taken in fixing smaller animals, as fluorescent protein expression can easily be disrupted.
In the method shown here, we describe an analysis of mosaic animals with GFP expressed in single tracheal cells. However, the same techniques are applicable to analysis of tracheal cells under a number of experimental manipulations. For instance, animals in which gene expression has been altered by molecular approaches, throughout the tracheal system or in individual cells by expression of RNAi transgenes or modified proteins, can also be examined with this approach. Non-genetic treatments which affect terminal cell development, such as drugs or hypoxia2,14, can also be characterized using these methods. In these latter cases, it is necessary to start with a Drosophila stock constitutively expressing GFP or another fluorescent protein throughout its tracheal system in order to visualize branching patterns. Gas-filling can be examined regardless of fluorescent protein expression.
Here, we only show examples of untagged (cytoplasmically localized) GFP being used to label the tracheal system. However, the methods described here are also suitable for detection of other native fluorescent proteins, such as DsRed, as well as fluorescent proteins which have been modified to be localized to specific subcellular structures. For instance, actin:GFP or tubulin:GFP fusions can be used to visualize the tracheal cytoskeleton10.

Cell shape is critical for function of individual cells within an organism, as well as cells that function as part of a tissue or organ. We use Drosophila tracheal terminal cells, a component of the insect respiratory system, to investigate the molecular mechanisms that participate in controlling two conserved types of cellular morphology: branching and tube formation (lumenogenesis). Terminal cells are located at the tips of a network of branched tubes that functions to deliver oxygen to internal tissues1 and have an elaborate branched morphology which depends on an FGF signaling pathway that is controlled by local oxygen levels within target tissues2. Terminal cell branches are thin tubes, with a gas-filled subcellular lumen running through each branch. The distinct cellular architectures of terminal cells, along with the ease by which genetic analysis can be performed in Drosophila, make these cells an excellent model for investigating mechanisms of cellular outgrowth, branching, and intracellular tube formation. Terminal cells have proved a useful model for understanding some of the signaling pathways leading to branched cell differentiation, outgrowth, and maturation2-4. Using this system unbiased, forward genetic screens for cell morphogenesis mutants have been performed, yielding insights into mechanisms controlling cell shape5,6. For instance, these screens have revealed that a specific RabGAP is required for cytoskeletal polarity and vesicle trafficking in lumen formation and positioning7; that integrin-mediated adhesion is required for branch stability8; and that epithelial PAR-polarity proteins regulate polarized membrane trafficking required for both branching and lumen formation9. Other studies in terminal cells have shown that asymmetric actin accumulation and microtubule organization is required for cell elongation and lumenogenesis10. Thus, diverse, conserved cell biological mechanisms contribute to terminal cell morphogenesis.
Here, we describe a method to rapidly fix intact third-instar Drosophila larvae for analysis of terminal cell branching and lumen formation. This protocol can also be carried out on both first and second instar animals. Key to this technique is the ability to visualize terminal cells that are genetically labeled by fluorescent protein expression directly through the larval cuticle of intact animals. Since this procedure does not require any post-fixation manipulations, such as antibody staining, to observe the cells, it is well suited to high throughput analysis, including genetic or drug screening. Fluorescent protein expression reveals the structure of the cytoplasmically-filled branches. Tube formation can be monitored in parallel using brightfield microscopy to identify the gas-filled lumen, which contrasts with the surrounding fluid-filled tissues.
Included in this protocol is the method for generating genetic mosaics based on the MARCM system11, to produce homozygous mutant terminal cells labeled with fluorescent proteins in otherwise unlabeled animals. This is necessary, since terminal cells only elaborate their complex structures relatively late in development; genetic mosaics allow for bypass of gene requirements in other tissues earlier in development. To generate MARCM clones, trachea are labeled using the tracheal-specific driver breathless (btl)12. Described here is the protocol for the Drosophila X chromosome; for other chromosomes, a similar procedure can be used, with genetic reagents appropriate to the chromosome being examined. Here, trachea are labeled by expression of a cytoplasmically localized GFP, but the procedure works equally well with expression of other fluorescent proteins, such as DsRed.
Additionally, we have included a method to quantify branching patterns and lumen formation in terminal cells, based on methods developed for characterizing neuronal branch patterns13. This kind of quantitative data can be critical in discerning the precise role of genes in the branching or lumenogenesis process, as well as allowing for direct comparisons between different mutants9.

<table><tbody><tr><td>Student Dumont #5 forceps</td><td>Fine Scientific Tools</td><td>91150-20</td><td></td></tr><tr><td>Leica MZ16 Dissecting microscope (or equivalent)</td><td>Leica</td><td>MZ16</td><td></td></tr><tr><td>AxioImager M1 compound microscope (or equivalent)</td><td>Carl Zeiss</td><td></td><td>Equiped with AxioCam MRm, A-PLAN 10X/0.25, EC PLAN-NEOFLUAR20X/0.5, Filter Set 38HE</td></tr><tr><td>100% Glycerol</td><td>BioExpress</td><td>M152-4L</td><td></td></tr><tr><td>Fly stock</td><td></td><td></td><td>Genotype: w FRT19A tub:GAL80 FLP122; Btl-GAL4, UAS-GFP y w FRT19A</td></tr><tr><td>Fly stock</td><td></td><td></td><td>Genotype: y w FRT19A</td></tr></tbody></table>

examination of drosophila larval tracheal terminal cells by light microscopy

Cell shape is critical for cell function. However, despite the importance of cell morphology, little is known about how individual cells generate specific shapes. Drosophila tracheal terminal cells have become a powerful genetic model to identify and elucidate the roles of genes required for generating cellular morphologies. Terminal cells are a component of a branched tubular network, the tracheal system that functions to supply oxygen to internal tissues. Terminal cells are an excellent model for investigating questions of cell shape as they possess two distinct cellular architectures. First, terminal cells have an elaborate branched morphology, similar to complex neurons; second, terminal cell branches are formed as thin tubes and contain a membrane-bound intracellular lumen. Quantitative analysis of terminal cell branch number, branch organization and individual branch shape, can be used to provide information about the role of specific genetic mechanisms in the making of a branched cell. Analysis of tube formation in these cells can reveal conserved mechanisms of tubulogenesis common to other tubular networks, such as the vertebrate vasculature. Here we describe techniques that can be used to rapidly fix, image, and analyze both branching patterns and tube formation in terminal cells within Drosophila larvae. These techniques can be used to analyze terminal cells in wild-type and mutant animals, or genetic mosaics. Because of the high efficiency of this protocol, it is also well suited for genetic, RNAi-based, or drug screens in the Drosophila tracheal system.

Results are shown in Figure 2. A single lateral group (LG) terminal cell shows extensive subcellular branching (visualized by GFP; A) and a gas-filled subcellular lumen running through each of the branches (visualized by brightfield microscopy; B). These images were collected from a mosaic L3 larva, generated using the MARCM system described in sections 1 &amp; 2, and heat fixed and imaged, as described in sections 3 &amp; 4. Panels C &amp; D show a NeuronJ generated trace, as described in section 5, of the branches and the lumen respectively of the images shown in A &amp; B. Panels E &amp; F show the location of the fat body (FB) branch in a larva prepared for imaging as described in sections 3 &amp; 4. Note that in this example, the animal is not a mosaic, and GFP is expressed throughout the entire tracheal system. Panels G and H show an example of a GFP-labeled terminal cell that was heat fixed for too long a time. GFP is diffuse (G), branches have broken down and portions of lumens are no longer gas filled, thus appearing as breaks in the brightfield image.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/50496/50496fig1highres.jpg" src="/files/ftp_upload/50496/50496fig1.jpg" /> 
 Figure 1. Larvae preparation and identification of terminal cells. (A) Place larva in a drop of 100% glycerol on a glass slide. (B) Multiple larvae can be placed for heat fixation. (C) After heat fixation, organize larvae parallel to each other and perpendicular to the long axis of the slide. (D) Place cover glass over larvae. (E) To reorient larvae, carefully push cover glass with forceps to roll the animals. (F) Wild-type third instar larva with GFP expressed throughout the tracheal system. The paired dorsal trunks (DT) are visible on the dorsal side. (G) The same larva after the rolling technique with the ventral side now facing upwards. (H) Diagram of lateral view of two third-instar tracheal hemisegments (one hemisegment is highlighted in grey). Circles indicate lateral group (LG) and fat body (FB) terminal cells which we use for quantitation. Dashed lines represent other branches of the tracheal system which we do not routinely quantitate. For a full description of the tracheal branches in a larval segment, refer to Ref 1.
<img alt="Figure 2" fo:content-width="3.5in" fo:src="/files/ftp_upload/50496/50496fig2highres.jpg" src="/files/ftp_upload/50496/50496fig2.jpg" /> 
 Figure 2. Representative images. (A-D) Mosaic L3 larvae were generated using the MARCM technique (section 1) and fixed and imaged using the protocol in sections 2-4. The branching pattern of a single LG terminal cell was visualized by mosaic expression of GFP (A); the gas-filled lumen visualized with brightfield microscopy (B). (C) Tracing of the branching pattern of the cell in A, generated using NeuronJ (section 5). (D) Tracing of the gas-filled lumen of the cell in B, generated using NeuronJ (protocol section 5). (E, F) A fat body (FB) terminal cell (highlighted by the red circle) visualized by GFP expression throughout the tracheal system (E) and brightfield microscopy (F) in a larva prepared by the protocol in sections 2-4. (G, H) Example of fixation artifacts obtained when the sample is heated for too long a period. GFP is diffuse and small branches have degraded (G). Areas of lumen also no longer appear air-filled (arrow in H). Scale bar: 100 &mu;m.

Watch this Scientific Journal Video about Examination of Drosophila Larval Tracheal Terminal Cells by Light Microscopy at JoVE.com

Examination of Drosophila Larval Tracheal Terminal Cells by Light Microscopy

Here, we present a method for light microscopy analysis of tracheal terminal cells in Drosophila larvae. This method allows for quick examination of branch and lumen morphology in whole animals and would be useful for analysis of individual mutants or screens for mutations affecting terminal cell development.

Examination of Drosophila Larval Tracheal Terminal Cells by Light Microscopy

Cell shape is critical for cell function. However, despite the importance of cell morphology, little is known about how individual cells generate specific ...

examination-drosophila-larval-tracheal-terminal-cells-light

Nanyang Technological University

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10.3K Views.  University of Utah. Here, we present a method for light microscopy analysis of tracheal terminal cells in Drosophila larvae. This method allows for quick examination of branch and lumen morphology in whole animals and would be useful for analysis of individual mutants or screens for mutations affecting terminal cell development.

Video: Examination of Drosophila Larval Tracheal Terminal Cells by Light Microscopy

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The Drosophila larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila is well known for the ease of powerful genetic manipulations and the larval nervous system has proven particularly useful in studying not only normal function but also perturbations that accompany some neurological disease (Lloyd and Taylor, 2010). Many key synaptic molecules found in Drosophila are also found in mammals and like most CNS excitatory synapses in mammals, the Drosophila NMJ is glutamatergic and demonstrates activity-dependent remodeling (Kohet al. , 2000). Additionally, Drosophila neurons can be individually identified because their innervation patterns are stereotyped and repetitive making it possible to study identified synaptic terminals, such as those between motor neurons and the body-wall muscle fibers that they innervate (Keshishian and Kim, 2004). The existence of evolutionarily conserved synapse components along with the ease of genetic and physical manipulation make the Drosophila model ideal for investigating the mechanisms underlying synaptic function (Budnik, 1996).
The active zones at synaptic terminals are of particular interest because these are the sites of neurotransmitter release. NC82 is a monoclonal antibody that recognizes the Drosophila protein Bruchpilot (Brp), a CAST1/ERC family member that is an important component of the active zone (Waghet al. , 2006). Brp was shown to directly shape the active zone T-bar and is responsible for effectively clustering Ca2+ channels beneath the T-bar density (Fouquetet al. , 2009). Mutants of Brp have reduced Ca2+ channel density, depressed evoked vesicle release, and altered short-term plasticity (Kittelet al. , 2006). Alterations to active zones have been observed in Drosophila disease models. For example, immunofluorescence using the NC82 antibody showed that the active zone density was decreased in models of amyotrophic lateral sclerosis and Pitt-Hopkins syndrome (Ratnaparkhiet al. , 2008; Zweieret al. , 2009). Thus, evaluation of active zones, or other synaptic proteins, in Drosophila larvae models of disease may provide a valuable initial clue to the presence of a synaptic defect.
Preparing whole-mount dissected Drosophila larvae for immunofluorescence analysis of the NMJ requires some skill, but can be accomplished by most scientists with a little practice. Presented is a method that provides for multiple larvae to be dissected and immunostained in the same dissection dish, limiting environmental differences between each genotype and providing sufficient animals for confidence in reproducibility and statistical analysis.

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The neuromuscular junction (NMJ) of Drosophila melanogaster is an important model system for studying normal synaptic function as well as perturbations to synaptic function found in certain neurological diseases. We present a protocol for dissection of the Drosophila larval motor system and immunostaining for active zone proteins within the NMJ.

The Drosophila  larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila  is well known for ...

Neuroscience

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

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

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

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

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

Developmental Biology

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval body wall contains several cell types that are models to study cell and tissue morphogenesis. For example tracheae are used to examine tube morphogenesis1, and the dendritic arborization (DA) sensory neurons of the Drosophila larva have become a primary system for the elucidation of general and neuron-class-specific mechanisms of dendritic differentiation2-5 and degeneration6.
The shape of dendrite branches can vary significantly between neuron classes, and even among different branches of a single neuron7,8. Genetic studies in DA neurons suggest that differential cytoskeletal organization can underlie morphological differences in dendritic branch shape4,9-11. We provide a robust immunological labeling method to assay in vivo microtubule organization in DA sensory neuron dendrite arbor (Figures 1, 2, Movie 1). This protocol illustrates the dissection and immunostaining of first instar larva, a stage when active sensory neuron dendrite outgrowth and branching organization is occurring 12,13.
In addition to staining sensory neurons, this method achieves robust labeling of microtubule organization in muscles (Movies 2, 3), trachea (Figure 3, Movie 3), and other body wall tissues. It is valuable for investigators wishing to analyze microtubule organization in situ in the body wall when investigating mechanisms that control tissue and cell shape.

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how complex cell shapes, such as neuronal dendrites, are achieved during development, it is important to be able to accurately assay microtubule organization. Here we describe a robust immunohistological labeling method to examine microtubule organization of dendritic arborization neuron sensory dendrites, trachea, muscle, and other Drosophila larva body wall tissues.

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval ...

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila larvae, and fish, which all engage in undulatory movements (like crawling or swimming) as their primary mode of locomotion. However, a more sophisticated understanding of the individual motor neuron (MN) specification&#8212;at least in terms of informing disease therapies&#8212;demands an equally tractable system that better models the complex appendage-based locomotion schemes of vertebrates. The adult Drosophila locomotor system in charge of walking meets all of these criteria with ease, since in this model it is possible to study the specification of a small number of easily distinguished leg MNs (approximately 50 MNs per leg) both using a vast array of powerful genetic tools, and in the physiological context of an appendage-based locomotion scheme. Here we describe a protocol to visualize the leg muscle innervation in an adult fly.

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Here we describe a protocol to visualize the axonal targeting with a florescent protein in adult legs of Drosophila by fixation, mounting, imaging, and post-imaging steps.

The majority of work on the neuronal specification has been carried out in genetically and physiologically tractable models such as C. elegans, Drosophila ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in communication with the epidermal cells they innervate and thus their analysis benefits from in situ visualization of both neuronally and epidermally expressed proteins by immunofluorescence. Traditional methods of preparing larval fillets for immunofluorescence experiments leave intact the muscle tissue that covers most of the body wall, presenting several challenges to imaging neuronal and epidermal proteins. Here we describe a method for removing muscle tissue from Drosophila larval fillets. This protocol enables imaging of proteins that are otherwise obscured by muscle tissue, improves signal to noise ratio, and facilitates the use of super-resolution microscopy to study da neuron development.

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Studies of neuronal morphogenesis using Drosophila larval dendritic arborization (da) neurons benefit from in situ visualization of neuronal and epidermal proteins by immunofluorescence. We describe a procedure that improves immunofluorescence analysis of da neurons and surrounding epidermal cells by removing muscle tissue from the larval body wall.

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in ...

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An ex vivo technique for imaging mouse airway epithelia for quantitative analysis of motile cilia function important for insight into mucociliary clearance function has been established. Freshly harvested mouse trachea is cut longitudinally through the trachealis muscle and mounted in a shallow walled chamber on a glass-bottomed dish. The trachea sample is positioned along its long axis to take advantage of the trachealis muscle to curl longitudinally. This allows imaging of ciliary motion in the profile view along the entire tracheal length. Videos at 200 frames/sec are obtained using differential interference contrast microscopy and a high speed digital camera to allow quantitative analysis of cilia beat frequency and ciliary waveform. With the addition of fluorescent beads during imaging, cilia generated fluid flow also can be determined. The protocol time spans approximately 30 min, with 5 min for chamber preparation, 5-10 min for sample mounting, and 10-15 min for videomicroscopy.

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An easy and reliable technique for visualizing and quantifying airway cilia motility and cilia generated flow using mouse trachea is described. This technique can be modified to determine how a wide range of factors influence cilia motility, including pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load.

An ex vivo technique for imaging mouse airway epithelia for quantitative  analysis of motile cilia function important for insight into mucociliary  ...

Drosophila Larva Imaginal Disc Dissection: A Method to Observe Developing Epithelia

Source: Morimoto, K. and Tamori, Y. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/55901/" target="_blank">Induction and Diagnosis of Tumors in Drosophila Imaginal Disc Epithelia</a>. J. Vis. Exp. (2017).
When staining Drosophila imaginal discs, often a rough dissection is performed where the discs are exposed but remain attached to the body. Once stained, the discs of interest are isolated and mounted. The example protocol demonstrates a procedure for wing imaginal disc dissection and mounting.

Source: Morimoto, K. and Tamori, Y. Induction and Diagnosis of Tumors in Drosophila Imaginal Disc Epithelia. J. Vis. Exp. (2017).
When staining Drosophila ...

JoVE Encyclopedia of Experiments

Drosophila melanogaster (fruit fly)

Larva

Larval Ring Gland Dissection: Isolation of Developmental Endocrine Organs from Drosophila

Source: Imura, E., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/55519/" target="_blank">Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila melanogaster</a>. J. Vis. Exp. (2017).
This video describes the Drosophila ring gland, a complex endocrine structure important for biosynthesis and secretion of ecdysteroids and other hormones. The featured protocol demonstrates its dissection, fixation, and immunostaining.

Source: Imura, E., et al. Protocols for Visualizing Steroidogenic Organs and Their Interactive Organs with Immunostaining in the Fruit Fly Drosophila ...

Examination of Drosophila Larval Tracheal Terminal Cells by Light Microscopy

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Chapters in this video

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

Visualize Drosophila Leg Motor Neuron Axons Through the Adult Cuticle

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

Drosophila Larva Imaginal Disc Dissection: A Method to Observe Developing Epithelia

Larval Ring Gland Dissection: Isolation of Developmental Endocrine Organs from Drosophila