Name: the drosophila imaginal disc tumor model: visualization and quantification of gene expression and tumor invasiveness using genetic mosaics
Uploaded: 2016-10-06T14:00:00.000+00:00
Duration: 10 min 31 s
Description: Watch this Scientific Journal Video about The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics at JoVE.com

We thank the Bloomington Stock Center (Bloomington, USA), Dirk Bohmann, Katja Br&#252;ckner and Istvan Ando for fly stocks, and antibodies. We thank Marek Jindra and Colin Donohoe for comments on the manuscript. This work was supported by the Sofja Kovalevskaja Award to M.U. from the Alexander von Humboldt Foundation and DFG project UH243/1-1 to M.U. from the German Research Foundation.

NOTE: This work utilizes the eyFLP1; act&#62;y+&#62;Gal4, UAS-GFP; FRT82B, tub-Gal80 MARCM 82B Green tester line 11. Crossing the MARCM 82B Green tester virgins to males of strains such as w; UAS-a; UAS-bRNAi, FRT82B cmut genotype will yield progeny in which mitotic recombination will occur between the right arms of the 3rd homologous chromosomes. In this way, clones homozygous mutant for gene c located distal to the FRT82B site will be generated within the EAD and brain neuroepithelium. These clones will express GFP, transgene a and dsRNA for gene b (Figure 1A). Various eyFLP-MARCM tester lines for recombination on the X, 2L, 2R, 3L and 3R have been established and are available within the fly community.
1. Fly Handling, Crosses, and Staging
<ol>
	<li>Expand the appropriate MARCM tester stock at room temperature by populating multiple bottles at the same time. Flip the adults into fresh bottles every 2-3 days so that enough virgins can be collected for subsequent crosses. 
	NOTE: When collecting virgins, avoid a prolonged exposure to CO2 as this compromises female fecundity.</li>
	<li>Depending on the scale of the experiment, set up fly crosses in vials using at least 10 MARCM tester virgins and 4 males (e.g., for imaging of fluorescent reporters and immunostaining) or in bottles with at least 30 virgins and 10 males (e.g., for RNA isolation and quantification of invasiveness).</li>
	<li>Raise progeny at 25 &#176;C under a long day 16 hr/8 hr light-dark cycle. Flip parents into new vials/bottles every 24 hr to facilitate reproducible staging of larvae and to prevent overcrowding. 
	NOTE: Starting on day 6 after egg laying (AEL), the late third instar control larvae stop feeding, start wandering and enter pupariation. In contrast, the onset of the larval-pupal transition can be delayed or non-existent in larvae with EAD bearing hyperplastic or malignant clonal tumors.</li>
</ol>
2. Collecting Third Instar Larvae
<ol>
	<li>For immunostaining, collect about 20 late third instar larvae (wandering on the wall and those of a comparable body size from the food). Use forceps to gently pick larvae from the vial or bottle and transfer them into an embryo glass dish filled with phosphate-buffered saline (PBS).
	<ol>
		<li>For RNA isolation and quantification of invasiveness, collect at least 80 late third instar larvae (wandering on the wall and those of a comparable body size from the food). Use a squirt bottle to squirt PBS into a fly bottle containing larvae until the surface is covered.</li>
		<li>Soften the top layers of food with a spatula and pour the food mash containing larvae into a Petri dish. Using forceps, gently pick larvae and transfer them into an embryo glass dish filled with PBS.</li>
	</ol>
	</li>
	<li>Wash larvae with PBS until all residual food is removed. Inspect the larvae under the fluorescent stereomicroscope with 8 to 16X magnification to discard all &#34;leopard larvae&#34; that carry random GFP-positive spots throughout the larval body (Figure 2A, 2B and see Discussion Section). 
	NOTE: For RNA isolation include a penultimate washing step in 70% EtOH for 1 min.</li>
	<li>Place the dish containing selected larvae in PBS on ice until dissection. Process larvae within 30 min to avoid adverse effects of cold and starvation.</li>
</ol>
3. Dissection of Larvae
<ol>
	<li>To dissect the EAD under a stereomicroscope, use one pair of forceps to gently press the middle part of the larva against the glass dish bottom. With the second forceps, grab the larval mouth hooks and pull them away from the body (Figure 2C). 
	NOTE: The pulling force is often sufficient to break the optic stalks connecting the EAD pair to the brain lobes. If the brain or its parts stay attached to EAD and their presence is not desired for further applications, cut the optic stalks with a help of the two pairs of forceps. For invasive tumors (e.g., rasV12scrib1) the connection between EAD and the brain lobes becomes increasingly obscured with time by overgrowing and migrating clonal cells.
	<ol>
		<li>To prepare the EAD/brain complex with an intact ventral nerve cord (VNC) (Figure 2D), use forceps to cut a larva in the middle of the body. Discard the posterior half.</li>
		<li>Hold the anterior half of the larval body between the tips of one forceps, flip the larva inside-out by pushing in the mouth hooks with the tip of the second forceps and by rolling the cuticle over it with the first forceps pair.</li>
	</ol>
	</li>
	<li>Using forceps carefully remove all extraneous tissues (e.g., salivary glands, fat body, and gut) while leaving the EAD pair or the EAD/brain complex connected to the mouth hooks. Disentangle the mouth hooks from the overlying cuticle.&#160;Free the VNC by severing the axonal projections extending to body muscles and epidermis (Figures 1B, 2C, 2D). 
	NOTE: The mouth hooks allow for safe manipulation of tissue with forceps and facilitate more efficient sinking and easy recognition of tissue during washing. Dissected EAD change morphology relatively fast when kept unfixed in PBS. Limit the dissection time to 20-30 min.</li>
	<li>To transfer the tissue, coat a P20 micropipette tip by pipetting the remaining body carcasses several times up and down. To transfer the larger EAD/brain complexes, cut the P20 micropipette tip with scissors. 
	NOTE: The coating procedure is critical to prevent sticking and loss of dissected tissue during transfer. Use of a P20 micropipette minimizes the transfer of PBS into fixative solution, limiting unwanted dilution.
	<ol>
		<li>For immunostaining, transfer the dissected EAD into a 0.5 ml tube filled with 400 &#181;l 4% paraformaldehyde (PFA) fixative. Use 1.5 ml tube with 1 ml PFA for larger EAD/brain complexes meant for scoring of invasiveness. 
		CAUTION: PFA is highly toxic. Wear protective gloves and clothing. Avoid contact with skin, eyes or mucous membranes.</li>
		<li>For RNA isolation transfer the dissected EAD into a new glass dish filled with cold PBS. Use forceps to clean EAD from the ring and lymph glands, to minimize sample contaminations.</li>
		<li>Detach the EAD from the mouth hooks by severing the connection between the antenna discs and mouth hooks using forceps (Figure 2E). To avoid RNA degradation and gene expression artifacts, limit the dissection time to 30 min. Proceed to step 7. 
		NOTE: An experienced researcher can dissect about 60 larvae in 30 min excluding time needed for larvae collection and washing. Total RNA yields depend on the EAD size that vary among genotypes and developmental stages. Dissection of 80-100 control EAD pairs from late third instar larvae should yield 5-8 &#181;g total RNA.</li>
	</ol>
	</li>
</ol>
4. Fixing, Immunostaining, and Mounting
<ol>
	<li>Fix samples in PFA fixative for 25 min while nutating at room temperature. The fixation time can be extended up to 60 min without altering tissue morphology and the antigenicity of proteins.</li>
	<li>Remove the fixative and wash samples with PBST (PBS with 0.1% Triton X-100) three times for 10 min using a nutator. Use sufficient volume of PBST for each washing step (400 &#181;l/0.5 ml tube). 
	NOTE: Fixed EAD/brain complexes meant for scoring of invasiveness do not require immunostaining. GFP signal remains well visible after fixation. Proceed to step 4.4 for the final dissection.
	<ol>
		<li>For immunostaining, block tissue in 200 &#181;l of a blocking solution (PBST with 0.3% BSA) for 20 min with gentle agitation at room temperature.
		<ol>
			<li>Incubate with primary antibodies diluted in 100 &#181;l of a blocking solution overnight at 4 &#176;C while gently shaking. Refer to primary antibody datasheet or published literature for recommended antibody dilution. 
			NOTE: The primary antibody dilution might need to be optimized.</li>
			<li>Following five 10 min washes in PBST, stain tissue with the fluorescent secondary antibodies diluted in blocking solution while gently shaking for 2 hr at room temperature or overnight at 4 &#176;C in the dark.</li>
			<li>Wash samples three times 10 min in PBST.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Replace PBST with 500 &#181;l of DAPI-staining solution. To label F-actin, add phalloidin conjugated to a fluorescent dye to DAPI-staining solution. After 15 min nutation in the dark, wash tissue once for 10 min with PBST. 
	NOTE: F-actin labeling can be carried out independently of DAPI staining.</li>
	<li>For the final dissection step, transfer tissue back into a PBS-filled glass dish using a 1 ml pipette. Using the two pairs of forceps clip off the mouth hooks.
	<ol>
		<li>Separate the brains that will be used for quantification of invasiveness from the EAD by cutting the optic stalk as overgrown EAD might hamper the analysis.</li>
	</ol>
	</li>
	<li>Place a drop of the mounting medium (15 &#181;l for a 22 mm x 22 mm coverslip) on an objective slide. With the help of forceps tips, distribute the medium into a thin layer. Transfer EAD, brains or the EAD/brain complexes into the mounting medium with a P20 micropipette.</li>
	<li>Use the forceps tips or a tungsten rod to distribute tissue and straighten the VNC on the slide.</li>
	<li>To avoid formation of bubbles while mounting, first touch one edge of a coverslip to the mounting medium and then slowly lower onto the mounting medium with the help of forceps. Use small pieces of filter paper or tissue wipe to absorb excess mounting medium around the coverslip edges. 
	NOTE: DABCO/Poly(vinyl alcohol) 4-88 mounting medium hardens at 4 &#176;C within an hour. Slides can be stored for months at 4 &#176;C.</li>
</ol>
5. Confocal Imaging
<ol>
	<li>Acquire single confocal sections and stacks using a confocal microscope equipped with 20X, 40X and 60X oil objectives.</li>
	<li>Prevent pixel saturation. Use same image acquisition parameters including image resolution, laser input power, gain, offset, frame averaging, a step size in a z-series and maintain these settings for all genotypes to allow for comparisons of pixel intensities between different genotypes 21,22. 
	NOTE: For visualization of EAD/brain complex, generate maximum projections and stitch respective, neighboring images. For final image preparation, use post image processing software for panel assembly and to adjust brightness and contrast.</li>
</ol>
6. Quantification of Tumor Invasiveness
<ol>
	<li>Define a scoring system that will characterize the different levels of tumor invasiveness considering the spatial pattern of invasion and the amount of GFP-positive cells spreading into the brain and VNC. Prepare evaluation sheets for documentation (Figure 3A).</li>
	<li>Mount at least 80 fixed intact brains for each genotype on one slide using 40 &#181;l of the mounting medium per 24 mm x 50 mm coverslip.
	<ol>
		<li>To allow unbiased scoring, ask a neutral party to anonymize the slides, so that the scoring procedure is unbiased. Evaluate anonymized slides by two lab members independently. Disclose genotypes only after the counting is finished.</li>
	</ol>
	</li>
	<li>Evaluate the degree of malignancy by a blind scoring of mounted brains under a fluorescent stereomicroscope equipped with a GFP filter set. Use a marker to label brains that have been already viewed to avoid double counting (Figure 3B).</li>
	<li>Calculate the percentage of brains falling into each of the defined invasive categories per genotype. Determine the statistical significance using a chi-square test (Figure 3C).</li>
</ol>
7. RNA Isolation, DNase Treatment, and Quality Check
NOTE: All reagents (solutions, plasticware) used in the following steps should be free of RNase activity. Always wear gloves and use a chemical hood for work with organic solvents.
<ol>
	<li>Transfer the clean EAD with a coated P20 micropipette tip into 1.5 ml tube.</li>
	<li>Let tissue sink to the bottom of the tube, carefully remove PBS and replace it with 100 &#181;l RNA lysis reagent (sufficient for up to 140 EAD).</li>
	<li>Lyse tissue by vortexing. At this point, samples can be deep frozen in liquid nitrogen and stored at -80 &#176;C or directly processed with a standard RNA isolation protocol. 
	NOTE: Samples of similar genotypes from multiple dissection rounds can be pooled once in RNA lysis reagent.</li>
	<li>Fill the sample volume to 0.9 ml with RNA lysis reagent. Add 0.2 ml of chloroform and mix samples vigorously for 15-20 sec.</li>
	<li>Following 2 to 3 min incubation at room temperature, centrifuge the samples at 10,000 x g for 15 min at 4 &#176;C.</li>
	<li>Transfer colorless upper aqueous phase containing RNA into a fresh tube without disturbing the interphase. Stay away from the interphase as it contains proteins and genomic DNA. 
	NOTE: The recovered volume should be about 60% of the volume of RNA lysis reagent used for homogenization.</li>
	<li>Precipitate the RNA by adding 0.5 ml of isopropyl alcohol. Vortex and incubate samples at room temperature for 10 min.</li>
	<li>Centrifuge the samples at 10,000 x g for 15 min at 4 &#176;C.</li>
	<li>Discard the supernatant and wash the RNA pellet once with 600 &#181;l of 75% ethanol. After a short vortex and centrifugation (10,000 x g for 7 min at 4 &#176;C) discard all ethanol and let the RNA pellet air-dry for 5-10 min placing the open tube on a clean tissue wipe. 
	NOTE: The RNA pellet might be visible as a tiny opaque/white stripe at the bottom of the tube or invisible. The remaining ethanol drops can be carefully removed with P10 micropipette tip.</li>
	<li>Dissolve RNA in 50 &#181;l of DEPC-treated water by vortexing or passing solution a few times through a pipette tip.</li>
	<li>To minimize contamination with genomic DNA, treat total RNA with DNase by adding 50 &#181;l of a mix containing 1 &#181;l of DNase (2 U/&#181;l) and 10 &#181;l of 10x DNase buffer in DEPC-treated water. Vortex and spin down.</li>
	<li>Incubate samples at 37 &#176;C in a water bath or a heating block for 30 to 40 min. After incubation, add 100 &#181;l of DEPC-treated water into each sample.</li>
	<li>To inactivate the enzymatic activity and clean RNA from DNase, add 200 &#181;l of phenol:chloroform:isoamyl alcohol (25:24:1) solution, vortex for 1 min and centrifuge the samples at 10,000 x g for 7 min at 4 &#176;C.</li>
	<li>Carefully transfer the upper aqueous phase containing RNA into a fresh tube. Add equal volume (200 &#181;l) of chloroform, mix and repeat the centrifugation step from above.</li>
	<li>Collect carefully the upper phase and precipitate RNA by adding 1/10 volume of 3 M sodium acetate, pH 5.2 prepared in DEPC H2O and 2.5 volumes of 100% ethanol. Mix thoroughly by vortexing. 
	NOTE: Adding 0.5 &#181;l glycogen (20 &#181;g/&#181;l) to a precipitation mix helps to visualize the RNA pellet. To minimize RNA loss, use siliconized, low-binding 1.5 ml microcentrifuge tubes.</li>
	<li>Precipitate RNA at -20 &#176;C for at least 1 hr. 
	NOTE: The incubation might be carried out overnight. Samples can be placed at -80 &#176;C as well.</li>
	<li>Pellet RNA by centrifugation at 10,000 x g for 30 min at 4 &#176;C. Wash and dry pellet following the description in 7.9.</li>
	<li>Dissolve RNA in 10-15 &#181;l of DEPC-treated water. Store RNA at -80 &#176;C.</li>
	<li>Determine quantity and purity of RNA by measuring OD at 230 nm, 260 nm and 280 nm using a spectrophotometer. Calculate RNA concentration by applying the convention that 1 OD at 260 nm equals 40 &#181;g/ml RNA. 
	NOTE: The A260/280 ratio of &#34;pure&#34; RNA equals 2.0 while an A260/230 ratio should be in the range of 2.0-2.2. RNA samples with an A260/280 ratio between 1.7 and 2.0 are suitable for downstream applications such as cDNA synthesis. For preparation of mRNA-seq libraries the quality and quantity of RNA should be checked using an automated electrophoresis system. The 28S/18S ratio should be above 1.8.</li>
</ol>

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The authors declare that they have no conflict of interest.

The techniques to generate genetic mosaics in Drosophila are among the most sophisticated tools for analyzing and manipulating gene function 33. The eyFLP-MARCM system has proven powerful and robust as it allows the induction of visually marked, genetically defined clones in a spatially restricted manner, i.e., in tissues where the eyeless enhancer is active 9,18. This is particularly important when multiple genetic lesions are combined in the same cells. While these highl...

Cancer represents one of the most genetically heterogeneous group of diseases, whose incidence and mortality is dramatically increasing, particularly among the elderly worldwide. Cancer originates clonally from a tumor-initiating cell that escapes inherent tumor suppressor mechanisms and divides out of control. The gradual accumulation of genetic lesions that cooperatively promote growth, proliferation and motility while inhibiting death and differentiation transforms the initial benign overgrowth into a highly malignant, metastatic and deadly tumor. It has become evident that in addition to genetic alterations, tumor progression requires changes in the surrounding stroma and crosstalk between the tumor and multiple cell types (e.g., fibroblasts, immune and endothelial cells) in its microenvironment. Understanding the molecular principles underlying malignant transformation including tumor-stroma interactions is of great importance for developing prevention and early screening strategies, as well as new and effective treatments to combat cancer metastasis and drug resistance.
The fruit fly Drosophila melanogaster has become an attractive system for cancer research 1-4 owing to its fast generation time, remarkable conservation of signaling nodes between flies and humans, limited genetic redundancy and wealth of advanced genetic tools that facilitate manipulation of almost any gene in a temporarily and spatially restricted manner. Genetically defined tumors of varying malignancy can be reproducibly engineered in Drosophila by introducing gain- and loss-of-function mutations in a subset of progenitors in an otherwise wild type tissue using the MARCM technique 5. The MARCM tool combines FLP/FRT (FLP recombinase/FLP Recognition Target)-mediated mitotic recombination 6 with FLP-out 7 and Gal4/UAS (Upstream Activation Sequence) 8 target gene expression systems 9. With this method expression of any UAS-based transgene, including oncogene or fluorescent protein cDNAs or inverted DNA repeats for dsRNA-induced gene silencing, will be restricted to a clone of cells that have lost a specific genetic locus and a Gal4 repressor due to recombination (Figure 1A). Clonal patches marked with green fluorescent (GFP) or red fluorescent proteins (e.g., RFP, DsRed, mCherry) can be easily tracked throughout development, isolated and analyzed. Importantly, their behavior can be directly compared to the adjacent wild type tissue. Thus, questions pertinent to the cell autonomous and non-autonomous effects of genetic lesions can be conveniently studied. Similar to mammals, only clones in which multiple oncogenic lesions are combined become malignant in Drosophila and recapitulate key hallmarks of mammalian cancer. They overproliferate, evade apoptosis, induce inflammation, become immortal and invasive, ultimately killing the host 10-17.
Here, we describe a protocol to generate genetically defined clonal tumors in the eye/antennal and brain tissue of Drosophila larvae using the MARCM technique. The method relies on a MARCM tester stock which expresses the yeast FLP recombinase under the control of the eyeless enhancer (eyFLP) 18,19. In this way, GFP-labeled clones are generated in both peripodial and columnar epithelium of the EAD and the neuroepithelium of the brain throughout embryonic and larval stages (Figure 1A, B and reference 20). Clones can be easily followed until adulthood as the EAD develops into the adult eye, antenna and head capsule while the neuroepithelium gives rise to neuroblasts that produce differentiated optic lobe neurons.
To facilitate extensive molecular, functional and phenotypic characterization of the mosaic tissue, we describe a protocol for dissection of the EAD and brain from the third instar larvae and outline how to process them for three different applications: (i) detection of transgenic fluorescent reporters and immunostaining, (ii) quantification of tumor invasiveness and (iii) analysis of gene expression changes using a quantitative real-time PCR (qRT-PCR) or a high-throughput mRNA sequencing (mRNA-seq) (Figure 1C).
The immunostaining protocol can be used to visualize any protein of interest with a specific antibody. Transgenic fluorescent transcriptional reporters provide convenient and precise spatiotemporal information on the activity of a particular signaling pathway. Cell-lineage specific reporters, on the other hand, reflect qualitative and quantitative changes in cell populations within the mosaic tissue and among tumors of distinct genotypes. Quantification of invasive behavior facilitates the comparison of tumor malignancy between genotypes. Finally, the protocol describing collection and processing of mosaic EAD for RNA isolation is suitable for both small and large-scale downstream applications such as reverse transcription followed by qRT-PCR and genome-wide mRNA-seq, respectively. The qualitative and quantitative data obtained from these assays provide novel insights into the social behavior of clonal tumors. Moreover, they produce a solid foundation for functional studies on the role of individual genes, genetic networks and tumor microenvironment in different stages and aspects of tumorigenesis.

<table><tbody><tr><td>Agar</td><td>Gew&amp;uuml;rzm&amp;uuml;hle Brecht, Eggenstein, Germany</td><td>00262-0500</td><td>Fly food recipe: Prepare 20 L fly food with 160 g agar, 360 g yeast, 200 g soy flour, 1.6 kg yellow cornmeal, 1.2 L malt extract, 300 ml light corn syrup, 130 ml propionic acid and 200 ml 15% nipagin. Fly food should be cooked for 1 hour at 85&amp;nbsp;&amp;deg;C.</td></tr><tr><td>Corn syrup</td><td>Grafschafter Krautfabrik Josef Schmitz KG, Meckenheim, Germany</td><td>01939</td><td></td></tr><tr><td>Propionic acid</td><td>Carl Roth GmbH, Karlsruhe, Germany</td><td>6026.1</td><td></td></tr><tr><td>Cornmeal</td><td>ReformKontor GmbH, Zarrentin, Germany</td><td>4010155063948</td><td></td></tr><tr><td>Malt extract</td><td>CSM Deutschland GmbH, Bremen, Germany</td><td>728985</td><td></td></tr><tr><td>Soy flour</td><td>Stockmeier Food GmbH, Herford, Germany</td><td>1000246441010</td><td></td></tr><tr><td>Yeast</td><td>Werner Ramspeck GmbH, Schwabach, Germany</td><td>210099K</td><td></td></tr><tr><td>Methyl-4-benzoate/ Nipagin</td><td>Sigma-Aldrich, Deisenhofen, Germany</td><td>H5501</td><td>Prepare a 15% stock solution with 70% EtOH</td></tr><tr><td>Drosophila fly food vials</td><td>Kisker Biotech, Steinfurt, Germany</td><td>789008</td><td></td></tr><tr><td>Vial plugs</td><td>K-TK e.K., Retzstadt, Germany</td><td>1002</td><td></td></tr><tr><td>Drosophila fly food bottles</td><td>Greiner Bio-One, Frickenhausen, Germany</td><td>960177</td><td></td></tr><tr><td>Bottle plugs</td><td>K-TK e.K., Retzstadt, Germany</td><td>1002 S</td><td></td></tr><tr><td>Poly(vinyl alcohol) 4-88/[-CH&lt;sub&gt;2&lt;/sub&gt;CHOH-]&lt;sub&gt;n&lt;/sub&gt;</td><td>Sigma-Aldrich, Deisenhofen, Germany</td><td>81381</td><td>Mounting medium recipe: Dissolve 6 g Poly(vinyl alcohol) 4-88 in 39&amp;nbsp;ml Millipore H&lt;sub&gt;2&lt;/sub&gt;O, 6 ml 1 M Tris (pH 8.5) and 12.5 ml glycerol. Stir overnight at 50&amp;nbsp;&amp;deg;C and centrifuge 20&amp;nbsp;min at 5,000 rpm. Add DABCO to the supernatant to get a final concentration of 2.5%. Store aliquots at -20&amp;nbsp;&amp;deg;C.</td></tr><tr><td>1,4-Diazabicyclo[2.2.2]octane/ DABCO&amp;nbsp;</td><td>Sigma-Aldrich, Deisenhofen, Germany</td><td>D2522</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich, Deisenhofen, Germany</td><td>9002-93-1</td><td></td></tr><tr><td>Phosphate-buffered saline/ PBS</td><td></td><td></td><td>137 mM NaCl, 2.7 mM KCl, 10 mM Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; and 1.8 mM KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, pH 7.4 in Millipore H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>PBST</td><td></td><td></td><td>0.1% Triton X-100 in PBS</td></tr><tr><td>Paraformaldehyde/ PFA</td><td>Sigma-Aldrich, Deisenhofen, Germany</td><td>158127</td><td>4% PFA fixative recipe: Dissolve 4&amp;nbsp;g PFA in 80 ml Millipore H&lt;sub&gt;2&lt;/sub&gt;O on a magnetic stirrer plate heated to 55&amp;nbsp;&amp;deg;C. Add 1 M NaOH dropwise until all PFA particles are dissolved. Add 10 ml 10x PBS and adjust the pH to 7.4 with 1 M HCl. Mix in 100 &amp;micro;l Triton X-100 and fill up with Millipore H&lt;sub&gt;2&lt;/sub&gt;O to 100 ml. Store aliquots at -20 &amp;deg;C. Avoid repeated thawing. (CAUTION: PFA is highly toxic. Prepare the PFA fixative in a fume hood. Wear a self-contained breathing apparatus, gloves and clothing. Avoid contact with skin, eyes or mucous membranes.)</td></tr><tr><td>Bovine Serum Albumin/ BSA</td><td>Sigma-Aldrich, Deisenhofen, Germany</td><td>A3059</td><td>Blocking solution recipe: Dissolve 0.3% BSA in PBST</td></tr><tr><td>Fluorescently labeled phalloidin</td><td>Invitrogen, Karlsruhe, Germany, Molecular Probes</td><td>A12379, A22283, A22284</td><td>Phalloidin stock solution: Dissolve powder (300 U) in 1.5 ml Methanol. Store at -20 &amp;deg;C. For staining, use in dilution 1:250 (A12379, A22283) and 1:100 (A22284) in PBST.</td></tr><tr><td>4&amp;rsquo;,6-Diamidino-2-phenylindol Dihydrochlorid/ DAPI</td><td>Carl Roth GmbH, Karlsruhe, Germany</td><td>6335</td><td>DAPI staining solution: Prepare a stock solution of 5 mg/ml in Millipore H&lt;sub&gt;2&lt;/sub&gt;O and store aliquots at 4 &amp;deg;C. Used in dilution 1:1,000 in PBST.</td></tr><tr><td>Mouse anti-H2 antibody</td><td>Kurucz et al., 2003</td><td></td><td>Used in dilution 1:500 in blocking solution</td></tr><tr><td>Cy5 AffiniPure Donkey Anti-Mouse IgG (H+L)</td><td>Jackson ImmunoResearch, Suffolk, UK</td><td>715-175-151</td><td>Used in dilution 1:500 in blocking solution</td></tr><tr><td>Dumont #5 forceps</td><td>Fine Science Tools, Heidelberg, Germany</td><td>11295-10</td><td></td></tr><tr><td>Glass embryo dish (30&amp;nbsp;mm)</td><td>Thermo Scientific, Schwerte, Germany</td><td>E90</td><td></td></tr><tr><td>Tungsten needles</td><td>Fine Science Tools, Heidelberg, Germany</td><td>10130-20</td><td></td></tr><tr><td>Nickel Plated Pin Holder</td><td>Fine Science Tools, Heidelberg, Germany</td><td>26018-17</td><td></td></tr><tr><td>Microscope slides</td><td>VWR, Darmstadt, Germany</td><td>631-1553</td><td></td></tr><tr><td>Coverslips 22 x 22 mm (#1 Menzel-Gl&amp;auml;ser)</td><td>VWR, Darmstadt, Germany</td><td>631-1336</td><td></td></tr><tr><td>Coverslips 24 x 50 mm (0.13-0.16&amp;nbsp;mm)</td><td>Thermo Scientific, Schwerte, Germany</td><td>1076371</td><td></td></tr><tr><td>Kimtech Science Precision Wipes</td><td>Thermo Scientific, Schwerte, Germany</td><td>06-677-70</td><td></td></tr><tr><td>Dissecting stereomicroscope</td><td>Olympus, Hamburg, Germany</td><td>SZX7 (DF PLAPO 1X-4 Japan)</td><td></td></tr><tr><td>Fluorescent stereomicroscope</td><td>Olympus, Hamburg, Germany</td><td>SZX16 (SDF PLAPO 0.8X Japan) with DP72 CCD camera</td><td>Equipped with the narrow blue bandpass filter set for excitation of GFP other blue excitable fluorochromes (excitation filter BP460-480 nm, barrier filter BA495-540 nm) and narrow green excitation and longpass barrier filter for RFP and other green excitable fluorochromes (excitation filter BP530-550, barrier filter BA575IF). Software: Olympus cellSens Standard 1.11.</td></tr><tr><td>Confocal microscope</td><td>Olympus, Hamburg, Germany</td><td>FV1000</td><td>Equipped with inverted IX81 microscope. Objectives: 20&amp;times; UPlan S-Apo (NA 0.85), 40&amp;times; UPlanFL (NA 1.30) and 60&amp;times; UPlanApo (NA 1.35). Lasers: UV laser Diode LD405 (50&amp;nbsp;mW), Argon laser, multi-line 457/(476)/ 488/515 (40&amp;nbsp;mW), Yellow/Green laser diode LD560 (15&amp;nbsp;mW) and Red Laser diode 635 (20&amp;nbsp;mW). Software: Fluoview 2.1c Software</td></tr><tr><td>Drosophila cooled incubator</td><td>Ewald Innovationstechnik GmbH, Bad Nenndorf, Germany</td><td>&amp;nbsp;Sanyo MIR553</td><td></td></tr><tr><td>Squirt bottle</td><td>VWR, Darmstadt, Germany</td><td>215-8105</td><td></td></tr><tr><td>BD Clay Adams Nutator Mixer</td><td>VWR, Darmstadt, Germany</td><td>15172-203</td><td></td></tr><tr><td>ELMI Digital Rocking Shaker</td><td>VWR, Darmstadt, Germany</td><td>DRS-12</td><td></td></tr><tr><td>5 PRIME Isol-RNA Lysis Reagent</td><td>VWR, Darmstadt, Germany</td><td>2302700</td><td>The protocol is compatible with other TRIzol-based reagents, &lt;em&gt;e.g.&lt;/em&gt;, TRIreagent from Sigma (Cat. Nr.T9424), TRIzol reagent from Thermofisher (Cat. Nr. 15596).</td></tr><tr><td>Diethyl pyrocarbonate/ DEPC</td><td>Sigma-Aldrich, Deisenhofen, Germany</td><td>D5758</td><td>Dilute DEPC 1:1,000 in Millipore H&lt;sub&gt;2&lt;/sub&gt;O, stir overnight and autoclave.</td></tr><tr><td>UltraPure Phenol:Chloroform:Isoamyl Alcohol (25:24:1)</td><td>ThermoFischer Scientific, Invitrogen, Karlsruhe, Germany</td><td>15593-031</td><td></td></tr><tr><td>Chloroform</td><td>Merck, Darmstadt, Germany</td><td>102445</td><td></td></tr><tr><td>TURBO DNase (2 U/&amp;micro;l)</td><td>Thermo Scientific, Schwerte, Germany</td><td>AM2238</td><td></td></tr><tr><td>Invitrogen UltraPure Glycogen</td><td>ThermoFischer Scientific, Invitrogen, Karlsruhe, Germany</td><td>10-814-010</td><td></td></tr><tr><td>Sodium acetate trihydrate</td><td>VWR, Darmstadt, Germany</td><td>27652.232</td><td>Prepare a 3 M Sodium acetate solution with DEPC-H&lt;sub&gt;2&lt;/sub&gt;O and adjust the pH to 5.2</td></tr><tr><td>2-Propanol</td><td>Merck, Darmstadt, Germany</td><td>109634</td><td></td></tr><tr><td>Ethanol</td><td>Merck, Darmstadt, Germany</td><td>100983</td><td>Prepare a 75% EtOH dilution with DEPC-H&lt;sub&gt;2&lt;/sub&gt;O.</td></tr><tr><td>UV/Vis-Spectrophotometre NanoDrop ND-8000</td><td>Thermo Scientific, Schwerte, Germany</td><td>ND-8000</td><td></td></tr><tr><td>Eppendorf Microcentrifuge (Refrigerated)</td><td>Thermo Scientific, Schwerte, Germany</td><td>5417R</td><td></td></tr><tr><td>Experion RNA StdSens Analysis Kit</td><td>Bio-Rad Laboratories GmbH, M&amp;uuml;nchen, Germany</td><td>7007103</td><td></td></tr><tr><td>Experion Automated Electrophoresis Station</td><td>Bio-Rad Laboratories GmbH, M&amp;uuml;nchen, Germany</td><td>7007010</td><td></td></tr><tr><td>SuperScript III Reverse Transcriptase</td><td>Thermo Scientific, Schwerte, Germany</td><td>18080044</td><td></td></tr><tr><td>Oligo d(T) Primer</td><td>Integrated DNA Technologies, Leuven, Belgium</td><td></td><td>Prepare 100 &amp;micro;M stock solutions in DEPC-H&lt;sub&gt;2&lt;/sub&gt;O and store at -20 &amp;deg;C</td></tr><tr><td>dNTP Mixture</td><td>Takara Bio Europe/Clonetech, Saint-Germain-en-Laye, France</td><td>4030</td><td>Store aliquots of 25 &amp;micro;l at -20&amp;deg;C</td></tr><tr><td>CFX96 Touch Real-Time PCR Detection System</td><td>Bio-Rad Laboratories GmbH, M&amp;uuml;nchen, Germany</td><td>1855195</td><td></td></tr><tr><td>iQ SYBR Green Supermix</td><td>Bio-Rad Laboratories GmbH, M&amp;uuml;nchen, Germany</td><td>170-8882</td><td></td></tr><tr><td>Hard-Shell PCR Plates 96-well, thin wall</td><td>Bio-Rad Laboratories GmbH, M&amp;uuml;nchen, Germany</td><td>HSP9601</td><td></td></tr><tr><td>Microseal &#039;B&#039; Film</td><td>Bio-Rad Laboratories GmbH, M&amp;uuml;nchen, Germany</td><td>MSB1001</td><td></td></tr><tr><td>&lt;em&gt;rp49&lt;/em&gt; Forward primer</td><td>Integrated DNA Technologies, Leuven, Belgium</td><td></td><td>5&#039; TCC TAC CAG CTT CAA GAT GAC 3&#039;</td></tr><tr><td>&lt;em&gt;rp49&lt;/em&gt; Reverse primer</td><td>Integrated DNA Technologies, Leuven, Belgium</td><td></td><td>5&#039; CAC GTT GTG CAC CAG GAA CT 3&#039;</td></tr><tr><td>&lt;em&gt;mmp1&lt;/em&gt; Forward primer</td><td>Integrated DNA Technologies, Leuven, Belgium</td><td></td><td>5&#039; AGG GCG ACA AGT ACT ACA AGC TGA 3&#039;</td></tr><tr><td>&lt;em&gt;mmp1&lt;/em&gt; Reverse primer</td><td>Integrated DNA Technologies, Leuven, Belgium</td><td></td><td>5&#039; ACG TCT TGC CGT TCT TGT AGG TGA 3&#039;</td></tr></tbody></table>

the drosophila imaginal disc tumor model: visualization and quantification of gene expression and tumor invasiveness using genetic mosaics

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the context of a whole organism. Sophisticated techniques to generate genetic mosaics facilitate induction of visually marked, genetically defined clones surrounded by normal tissue. The clones can be analyzed through diverse molecular, cellular and omics approaches. This study describes how to generate fluorescently labeled clonal tumors of varying malignancy in the eye/antennal imaginal discs (EAD) of Drosophila larvae using the Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. It describes procedures how to recover the mosaic EAD and brain from the larvae and how to process them for simultaneous imaging of fluorescent transgenic reporters and antibody staining. To facilitate molecular characterization of the mosaic tissue, we describe a protocol for isolation of total RNA from the EAD. The dissection procedure is suitable to recover EAD and brains from any larval stage. The fixation and staining protocol for imaginal discs works with a number of transgenic reporters and antibodies that recognize Drosophila proteins. The protocol for RNA isolation can be applied to various larval organs, whole larvae, and adult flies. Total RNA can be used for profiling of gene expression changes using candidate or genome-wide approaches. Finally, we detail a method for quantifying invasiveness of the clonal tumors. Although this method has limited use, its underlying concept is broadly applicable to other quantitative studies where cognitive bias must be avoided.

To demonstrate the potential of the eyFLP-MARCM technique to generate GFP-marked patches of defined genotypes in Drosophila EAD, three types of clones were induced: (1) control expressing GFP only, (2) malignant tumors expressing an oncogenic form of the small G-protein Ras (RasV12) in a background of homozygous loss of a tumor suppressor gene scribble (scrib1), and (3) overgrowing but non-invasive rasV12scrib1jnk<...

Watch this Scientific Journal Video about The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics at JoVE.com

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

This protocol demonstrates how to generate fluorescently marked, genetically defined clonal tumors in the Drosophila eye/antennal imaginal discs (EAD). It describes how to dissect the EAD and brain from the third instar larvae and how to process them to visualize and quantify gene expression changes and tumor invasiveness.

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

Drosophila melanogaster has emerged as a powerful experimental system for functional and mechanistic studies of tumor development and progression in the ...

the-drosophila-imaginal-disc-tumor-model-visualization-quantification

Research

JoVE Journal

Cancer Research

11.0K Views.  University of Cologne.  Dit protocol laat zien hoe fluorescent gemerkt, genetisch bepaald klonale tumoren in de Drosophila oog / antennaal verbeelding schijven (EAD) te genereren. Het beschrijft hoe de EAD en de hersenen ontleden van de derde instar larven en hoe ze te verwerken te visualiseren en te kwantificeren genexpressie veranderingen en tumor invasiviteit. 

Video: The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

Mosaic Zebrafish Transgenesis for Functional Genomic Analysis of Candidate Cooperative Genes in Tumor Pathogenesis

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers. To robustly and efficiently identify oncogenic &#8220;drivers&#8221; among these tumors and define their complex relationships with concurrent genetic alterations during tumor pathogenesis remains a daunting task. Recently, zebrafish have emerged as an important animal model for studying human diseases, largely because of their ease of maintenance, high fecundity, obvious advantages for in vivo imaging, high conservation of oncogenes and their molecular pathways, susceptibility to tumorigenesis and, most importantly, the availability of transgenic techniques suitable for use in the fish. Transgenic zebrafish models of cancer have been widely used to dissect oncogenic pathways in diverse tumor types. However, developing a stable transgenic fish model is both tedious and time-consuming, and it is even more difficult and more time-consuming to dissect the cooperation of multiple genes in disease pathogenesis using this approach, which requires the generation of multiple transgenic lines with overexpression of the individual genes of interest followed by complicated breeding of these stable transgenic lines. Hence, use of a mosaic transient transgenic approach in zebrafish offers unique advantages for functional genomic analysis in vivo. Briefly, candidate transgenes can be coinjected into one-cell-stage wild-type or transgenic zebrafish embryos and allowed to integrate together into each somatic cell in a mosaic pattern that leads to mixed genotypes in the same primarily injected animal. This permits one to investigate in a faster and less expensive manner whether and how the candidate genes can collaborate with each other to drive tumorigenesis. By transient overexpression of activated ALK in the transgenic fish overexpressing MYCN, we demonstrate here the cooperation of these two oncogenes in the pathogenesis of a pediatric cancer, neuroblastoma that has resisted most forms of contemporary treatment.

The goal of this study is to demonstrate how the mosaic transgenesis strategy can be used in zebrafish to rapidly and efficiently assess the relative contributions of multiple oncogenes in tumor initiation and progression in vivo.

Comprehensive genomic analysis has uncovered surprisingly large numbers of genetic alterations in various types of cancers.  To robustly and efficiently ...

Developmental Biology

Induction and Diagnosis of Tumors in Drosophila Imaginal Disc Epithelia

In the early stages of cancer, transformed mutant cells show cytological abnormalities, begin uncontrolled overgrowth, and progressively disrupt tissue organization. Drosophila melanogaster has emerged as a popular experimental model system in cancer biology to study the genetic and cellular mechanisms of tumorigenesis. In particular, genetic tools for Drosophila imaginal discs (developing epithelia in larvae) enable the creation of transformed pro-tumor cells within a normal epithelial tissue, a situation similar to the initial stages of human cancer. A recent study of tumorigenesis in Drosophila wing imaginal discs, however, showed that tumor initiation depends on the tissue-intrinsic cytoarchitecture and the local microenvironment, suggesting that it is important to consider the region-specific susceptibility to tumorigenic stimuli in evaluating tumor phenotypes in imaginal discs. To facilitate phenotypic analysis of tumor progression in imaginal discs, here we describe a protocol for genetic experiments using the GAL4-UAS system to induce neoplastic tumors in wing imaginal discs. We further introduce a diagnosis method to classify the phenotypes of clonal lesions induced in imaginal epithelia, as a clear classification method to discriminate various stages of tumor progression (such as hyperplasia, dysplasia, or neoplasia) had not been described before. These methods might be broadly applicable to the clonal analysis of tumor phenotypes in various organs in Drosophila.

Induction and Diagnosis of Tumors in Drosophila Imaginal Disc Epithelia

Mosaic clone analysis in Drosophila imaginal disc epithelia is a powerful model system to study the genetic and cellular mechanisms of tumorigenesis. Here we describe a protocol to induce tumors in Drosophila wing imaginal discs using the GAL4-UAS system, and introduce a diagnosis method to classify the tumor phenotypes.

In the early stages of cancer, transformed mutant cells show cytological abnormalities, begin uncontrolled overgrowth, and progressively disrupt tissue ...

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Neuroscience

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 ...

A Protocol for Genetic Induction and Visualization of Benign and Invasive Tumors in Cephalic Complexes of Drosophila melanogaster

Drosophila has illuminated our understanding of the genetic basis of normal development and disease for the past several decades and today it continues to contribute immensely to our understanding of complex diseases 1-7. Progression of tumors from a benign to a metastatic state is a complex event 8 and has been modeled in Drosophila to help us better understand the genetic basis of this disease 9. Here I present a simple protocol to genetically induce, observe and then analyze the progression of tumors in Drosophila larvae. The tumor induction technique is based on the MARCM system 10 and exploits the cooperation between an activated oncogene, RasV12 and loss of cell polarity genes (scribbled, discs large and lethal giant larvae) to generate invasive tumors 9. I demonstrate how these tumors can be visualized in the intact larvae and then how these can be dissected out for further analysis. The simplified protocol presented here should make it possible for this technique to be utilized by investigators interested in understanding the role of a gene in tumor invasion.

A Protocol for Genetic Induction and Visualization of Benign and Invasive Tumors in Cephalic Complexes of Drosophila melanogaster

Cooperation between an activated oncogene like RASV12 and mutations in cell polarity genes like scribbled, result in tumor growth in Drosophila where tumor cells also display invasive behaviors. Here a simple protocol for the induction and observation of the benign and invasive tumors is presented.

Drosophila has illuminated our  understanding of the genetic basis of normal development and disease for the  past several decades and today it continues ...

Medicine

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

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

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

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

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

Biology

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

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

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

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

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

Laser Microdissection Applied to Gene Expression Profiling of Subset of Cells from the Drosophila Wing Disc

Heterogeneous nature of tissues has proven to be a limiting factor in the amount of information that can be generated from biological samples, compromising downstream analyses. Considering the complex and dynamic cellular associations existing within many tissues, in order to recapitulate the in vivo interactions thorough molecular analysis one must be able to analyze specific cell populations within their native context. Laser-mediated microdissection can achieve this goal, allowing unambiguous identification and successful harvest of cells of interest under direct microscopic visualization while maintaining molecular integrity. We have applied this technology to analyse gene expression within defined areas of the developing Drosophila wing disc, which represents an advantageous model system to study growth control, cell differentiation and organogenesis. Larval imaginal discs are precociously subdivided into anterior and posterior, dorsal and ventral compartments by lineage restriction boundaries. Making use of the inducible GAL4-UAS binary expression system, each of these compartments can be specifically labelled in transgenic flies expressing an UAS-GFP transgene under the control of the appropriate GAL4-driver construct. In the transgenic discs, gene expression profiling of discrete subsets of cells can precisely be determined after laser-mediated microdissection, using the fluorescent GFP signal to guide laser cut. 
Among the variety of downstream applications, we focused on RNA transcript profiling after localised RNA interference (RNAi). With the advent of RNAi technology, GFP labelling can be coupled with localised knockdown of a given gene, allowing to determinate the transcriptional response of a discrete cell population to the specific gene silencing. To validate this approach, we dissected equivalent areas of the disc from the posterior (labelled by GFP expression), and the anterior (unlabelled) compartment upon regional silencing in the P compartment of an otherwise ubiquitously expressed gene. RNA was extracted from microdissected silenced and unsilenced areas and comparative gene expression profiling determined by quantitative real-time RT-PCR. We show that this method can effectively be applied for accurate transcriptomics of subsets of cells within the Drosophila imaginal discs. Indeed, while massive disc preparation as source of RNA generally assumes cell homogeneity, it is well known that transcriptional expression can vary greatly within these structures in consequence of positional information. Using localized fluorescent GFP signal to guide laser cut, more accurate transcriptional analyses can be performed and profitably applied to disparate applications, including transcript profiling of distinct cell lineages within their native context.

Laser Microdissection Applied to Gene Expression Profiling of Subset of Cells from the Drosophila Wing Disc

Laser microdissection was applied to analyse gene expression profiling in specific compartments of Drosophila wing disc subjected to localised RNAi in vivo. RNA extracted from equivalent areas of silenced and unsilenced compartments was analysed by quantitative RT-PCR to determine comparative gene expression profiling within the context of native tissue microecology.

Heterogeneous nature of tissues has proven to be a limiting factor in the amount of information that can be generated from biological samples, ...

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called imaginal discs. These discs give rise to a high percentage of adult structures that are found within the adult fly. Here we describe a protocol that has been optimized to recover these discs and prepare them for analysis with antibodies, transcriptional reporters and protein traps. This procedure is best suited for thin tissues like imaginal discs, but can be easily modified for use with thicker tissues such as the larval brain and adult ovary. The written protocol and accompanying video will guide the reader/viewer through the dissection of third instar larvae, fixation of tissue, and treatment of imaginal discs with antibodies. The protocol can be used to dissect imaginal discs from younger first and second instar larvae as well. The advantage of this protocol is that it is relatively short and it has been optimized for the high quality preservation of the dissected tissue. Another advantage is that the fixation procedure that is employed works well with the overwhelming number of antibodies that recognize Drosophila proteins. In our experience, there is a very small number of sensitive antibodies that do not work well with this procedure. In these situations, the remedy appears to be to use an alternate fixation cocktail while continuing to follow the guidelines that we have set forth for the dissection steps and antibody incubations.

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

The adult structures of Drosophila are derived from sac-like structures called imaginal discs. Analysis of these discs provides insight into many developmental processes including tissue determination, compartment boundary establishment, cell proliferation, cell fate specification, and planar cell polarity. This protocol is used to prepare imaginal discs for light/fluorescent microscopy.

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called ...

Dissection and Mounting of Drosophila Pupal Eye Discs

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes. It contains approximately 800 separate eye units, termed ommatidia1. Each ommatidium contains eight neuronal photoreceptors that develop from undifferentiated cells following the passage of the morphogenetic furrow in the third larval instar2. Following the sequential differentiation of the photoreceptors, non-neuronal cells develop, including cone and pigment cells, along with mechanosensory bristle cells3. Final differentiation processes, including the structured arrangement of all the ommatidial cell types, programmed cell death of undifferentiated cell types and rhodopsin expression, occurs through the pupal phase4-7. This technique focuses on manipulating the pupal eye disc, providing insight and instruction on how to dissect the eye disc during the pupal phase, which is inherently more difficult to perform than the commonly dissected third instar eye disc. This technique also provides details on immunostaining to allow the visualization of various proteins and other cell components.

Dissection and Mounting of Drosophila Pupal Eye Discs

The goal of this technique is to enable researchers to perform dissection, immunostaining and mounting of pupal eye discs from Drosophila melanogaster of any age.

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes.  It contains approximately 800 ...

The Drosophila Imaginal Disc Tumor Model: Visualization and Quantification of Gene Expression and Tumor Invasiveness Using Genetic Mosaics

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