Work conducted in the L&eacute;cuyer laboratory is supported by funding from the National Sciences and Engineering Council of Canada (NSERC), the Canadian Institutes of Health Research (CIHR) and the Fonds de Recherche en Sant&eacute; du Qu&eacute;bec (FRSQ). Fabio Alexis Lefebvre and Ga&euml;l Moquin-Beaudry are supported by NSERC undergraduate research studentships, while Carole Iampietro is supported by the Angelo Pizzagalli postdoctoral fellowship.

1. RNA Probe Preparation
Overview: The following section describes the steps required to make Digoxigenin (Dig)-labeled RNA probes suitable for FISH. The first step involves cloning or PCR amplifying a sequence corresponding to the transcribed region of a gene of interest that will be used to generate a sequence-specific probe. This can be achieved by first cloning the gene segment into a plasmid in which the multiple cloning site is flanked by bacteriophage promoter elements (T7, T3 or Sp6), which can then serve as a template for PCR using flanking primers that incorporate the promoter sequences, thus generating a linear PCR product with different promoter sequences at each extremity (Figure 1A). Alternatively, to avoid the cloning step, one can also perform PCR amplification from genomic DNA or cDNA using oligonucleotides that include T7, T3 or Sp6 sequences at their 5' extremities (e.g. T7: 5'-TAATACGACTCACTATAGGGAGA-3'; T3: 5'-AATTAACCCTCACTAAAGGGAGA-3'; Sp6: 5'-ATTTAGGTGACACTATAGAAGAG-3'). The linear PCR products are then used as templates to make Dig-labeled sense or antisense RNA probes by run-off in vitro transcription with the appropriate polymerase (Figure 1B). When making probes for specific genes, we recommend preparing both sense and antisense RNA probes using distinct polymerases, which will be helpful for assessing FISH signal specificity (i.e. unless the gene of interest is transcribed in the antisense orientation, the sense RNA probe will reveal the level of background signal). In all of the following steps, one should take great care to avoid potential degradation by contaminating ribonucleases (RNAses) by first cleaning all bench areas and equipment with 70% ethanol or commercial RNAse decontamination solutions and by using RNAse-free supplies (e.g. water, tips, microcentrifuge tubes).
Materials
<ul>
 <li>1.5 ml microcentrifuge tubes, (Abgene, Rochester, NY, USA Cat. No 0900)</li>
 <li>Gel-extraction kits (QIAGEN, Mississauga, ON, Canada; Cat. No. 28706).</li>
 <li>RNAse free water (Wisent, Inc. Cat. No. 809-115-CL)</li>
 <li>RNA polymerases (T7, T3, or SP6). (20 U/ &mu;l, Fermentas Biosciences. Cat. No. EP0101, EP0111, EP0131).</li>
 <li>RNAse-OUT Ribonuclease inhibitors (40 U/ &mu;l), (Invitrogen, Burlington ON. Canada.Cat. No. 10777-019).</li>
 <li>Digoxigenin (DIG) nucleotide mixes (DIG-11-UTP, Roche Applied Biosciences, Laval, QC, Canada. Cat. No.11209256910).</li>
</ul>
Equipment
<ul>
 <li>Table top micro-centrifuge</li>
 <li>Gel-electrophoresis apparatus</li>
</ul>
<ol>
 <li>PCR amplify gene-specific sequence from genomic DNA, cDNA, or plasmid DNA to generate a linear PCR product flanked by bacteriophage T7, T3 or Sp6 promoter sequences. Follow the manufacturer's recommendations for PCR conditions.</li>
 <li>Verify the quality and size of the PCR product by agarose gel electrophoresis. Excise and purify PCR product using a standard gel-extraction kit according to the manufactures recommendations and elute the product in 50 &mu;l of buffer.</li>
 <li>Precipitate the PCR product by adding 5 &mu;l of 3M sodium acetate (pH 5.2) and 150 &mu;l of ice cold 100% ethanol, and place the tubes at -70 &deg;C overnight. Centrifuge tubes at 13,000 x g for 10 min at 4 &deg;C and wash the pellet with 70% ethanol. Spin again, remove all traces of ethanol and air dry the pellet. Resuspend in 25-50 &mu;l of RNAse-free water.</li>
 <li>Transcribe Dig-labeled probe using 200-500 ng purified PCR product in a 20 &mu;l reaction as follows: 2 &mu;l of 10X T7 transcription buffer (Invitrogen), 1 &mu;l (20 U/ &mu;l) of RNAse inhibitor, 2 &mu;l Dig-NTP mix, and 2 &mu;l T7 RNA polymerase (20 U/ &mu;l). Bring the final volume to 20 &mu;l with RNAse-free water. Incubate the transcription reaction at 37 &deg;C for 3-4 hr.</li>
 <li>Adjust transcription mix to 50 &mu;l with RNAse-free water and precipitate the Dig-labeled RNA probe as described in Step 4. Resuspend the washed RNA pellet in 100 &mu;l of RNAse-free water. Store the resuspended samples at -70 &deg;C.</li>
 <li>Quantify the probe using a nanodrop spectrophotometer. To verify the probe quality, run a 1-2 &mu;l sample on a 1-2% agarose gel stained with ethidium bromide.</li>
</ol>
2. Collection and Fixation of Drosophila Embryos
Overview: This section describes steps for harvesting and processing staged Drosophila embryo. Depending on the number of embryos needed, flies can be maintained in population cages of various sizes. The following steps are for collections performed using 900 cm3 cylindrical cages using 100mm apple juice collection plates. Proper fixation requires the removal of two protective layers surrounding the embryo: the outer chorion and the inner vitelline membrane 13. Once harvested, the embryos are first bathed in a 50% bleach solution to remove the chorion, then they are mixed in a biphasic solution containing heptane (permeabilizes the vitelline membrane), and PBS containing 3.7% formaldehyde. The vitelline membrane is then cracked and removed by transferring the embryos into a biphasic mixture of heptane and methanol. Proper embryo fixation and removal of the vitelline membrane is critical for the success of the downstream FISH procedure.
Materials
<ul>
 <li>Apple-juice agar plates (40% apple juice, 4% sucrose, 3.5% agar, 0.3% methyl paraben) with dime-sized yeast paste (baker's yeast mixed with water to butter consistency)</li>
 <li>Powdered paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA, USA; Cat. No. 19208)</li>
 <li>Heptane</li>
 <li>Methanol</li>
 <li>3% Bleach solution</li>
 <li>20 ml glass scintillation vials (Thermo Fisher Scientific, Ottawa, ON, Canada; Cat. No. 03-337-15).</li>
</ul>
Equipment
<ul>
 <li>Population cages</li>
 <li>Collection baskets (made using a Nitex mesh and the top portion of a cut 50 ml polypropylene tube in which a hole has been cut in the lid).</li>
 <li>Small paint brush</li>
 <li>Table top vortex</li>
</ul>
<ol>
 <li>Pre-clear well-fed population cages with a fresh apple juice/yeast plate for 1 hr at 25 &deg;C.</li>
 <li>Add a new apple juice/yeast plate to the cage to harvest early stage embryos. Incubate the cage for 4 hr at 25 &deg;C (note that collection times can be adjusted depending on the desired stages).</li>
 <li>Prepare a fresh 37% formaldehyde solution by combining 3.7 g paraformaldehyde powder, 70 &mu;l of 2N KOH, and 7.3 ml of milliQ water in a glass scintillation vial under a chemical hood. Boil the solution until the paraformaldehyde powder is completely dissolved, then cool to room temperature and filter into new scintillation vial.</li>
 <li>Prepare a biphasic fixation solution by combining 2.25 ml of 1X PBS with 250 ml of 37% formaldehyde in a scintillation vial, then adding 7.5 ml of heptane.</li>
 <li>Harvest the apple juice plate from the cage and collect the embryos by adding a bit of room temperature tap water and delicately brushing the embryos off the plate with the paint brush. Transfer the resuspended embryos into a basket and rinse with lukewarm tap water.</li>
 <li>Dechorionate embryos for 90 sec by bathing the collection baskets in bleach solution, then rinse thoroughly with lukewarm tap water (until bleach smell is gone).</li>
 <li>Disassemble the collection basket, collect embryos gently using a paintbrush and transfer them into the biphasic fixation solution. The embryos will accumulate at the interface between the PBS and heptane layers.</li>
 <li>Seal scintillation vials and fasten to a vortex mixer. Shake for 20 min on lowest setting.</li>
 <li>Remove and discard most of the lower PBS and upper heptane phases using a Pasteur pipette. Aspirate the remaining PBS/heptane/embryos mixture into the pipette. In a dropwise fashion, discard the lower PBS phase from the pipette, taking care not to eliminate the embryos. Deposit the embryos into a 1.5 ml tube containing a biphasic solution of 500 &mu;l heptane (upper phase) and 500 &mu;l methanol (lower phase).</li>
 <li>Shake tubes vigorously by hand for 30-45 sec to crack the vitelline membranes. Once cracked, the embryos will sink into the methanol.</li>
 <li>Discard the upper heptane phase and uncracked embryos at the heptane/methanol interphase, and add 500 &mu;l of methanol. Shake for 5 sec.</li>
 <li>Wash 3 times with methanol and store the embryos at -20 &deg;C (at this point embryos can be stored for weeks/months).</li>
</ol>
3. Post-fixation Processing, Hybridization and Post-hybridization Washes
Overview: This section details the processing steps for embryo permeabilization prior to FISH, pre-hybridization, hybridization with the RNA probes and post-hybridization washes. For the initial permeabilization steps the embryos are handled as a pool in a single tube (steps 1-9 below, aiming for 10-15 &mu;l of settled embryos/sample), but they are aliquoted into individual wells of a 96-well PCR plate before proceeding with the pre-hybridization step (step 10 below). This helps to ensure even treatment of all embryos in the initial phases of the experiment. When setting up a FISH experiment, it is important to include negative controls to determine the level of background signal in individual experiments. For this, we recommend including a 'no-probe' sample and, as noted in section 1, a sample hybridized with the 'sense' RNA probe, which will typically give a signal comparable to the 'no-probe' condition.
Materials:
<ul>
 <li>Methanol</li>
 <li>Phosphate buffered saline with 0.1% Tween-20 (PBT)</li>
 <li>Proteinase-K 1 mg/ml stock solution (Sigma Aldrich, Oakville, ON, Canada. Catalog No. P2308)</li>
 <li>Glycine (20 mg/ml stock solution)</li>
 <li>Hybridization solution: 50% Formamide, 5X SSC, 0.1% Tween-20, 100 g/ml of sheared salmon sperm DNA, 100 &mu;g/ml Heparin.</li>
 <li>0.2 ml halfskirted 96-well PCR plates, Abgene, Rochester, NY, USA Cat. No 0900)</li>
</ul>
Equipment
<ul>
 <li>Hybridization oven, digital heating block or water bath adjustable to 56 &deg;C, 80 &deg;C or 100 &deg;C, or a PCR machine.</li>
 <li>Multichannel pipettors (recommended to simplify washing steps)</li>
 <li>Nutating mixer</li>
</ul>
<ol>
 <li>Rinse the embryos in methanol, then in a 1:1 mixture of methanol:PBT, and then twice in PBT.</li>
 <li>Fix the embryos for 20 min in 1 ml of PBT containing 3.7% formaldehyde (freshly prepared as described in step 11) with constant rocking on a nutator.</li>
 <li>Wash embryos for 3 times for 2 min in PBT while rocking.</li>
 <li>Prepare a solution of 3 &mu;g/ml of proteinase K (PK) diluted in PBT and add 500 &mu;l of solution to samples. Incubate samples for 10 min at room temperature without shaking. Mix embryos every 2 min by jetting with a pipette or by gently inverting the tubes.</li>
 <li>Transfer embryos samples on ice for 1 hr.</li>
 <li>Move samples to room temperature and remove PK solution. Wash 2 times for 2 min with 1 ml of PBT + 2 mg/ml glycine.</li>
 <li>Fix samples 20 min in 1 ml of PBT + 3.7% formaldehyde with rocking. Rinse embryos 5 times in PBT.</li>
 <li>Rinse embryos with 1 ml of a 1:1 mixture of PBT and Hyb solution. Replace with 0.5-1 ml of 100% Hyb solution (at this step the embryos can be stored for several days/weeks at -20 &deg;C).</li>
 <li>Aliquot embryos into individual wells of a 96-well plate (aim for ~10-15 &mu;l of settled embryos/well, take care to use wide aperture tips to avoid damaging the embryos).</li>
 <li>Boil 100 &mu;l/sample of Hyb solution for 5 min and then cool on ice for 5 min. This solution is used for sample pre-hybridization (Pre-Hyb).</li>
 <li>Add 100 &mu;l/sample of Pre-Hyb solution and incubate for at least 2 hr at 56 &deg;C.</li>
 <li>For each sample, dilute ~100 ng of probe in 100 &mu;l of Hyb solution. Heat at 80 &deg;C for 3 min, then cool on ice for 5 min (heated probes can be kept on ice for several hours).</li>
 <li>Remove the Pre-Hyb solution from the embryos and replace with the RNA probe solution.</li>
 <li>Hybridize at 56 &deg;C for 12-16 hr.</li>
 <li>Pre-warm the following wash solutions at 56 &deg;C: (A) 200 &mu;l/sample of Hyb solution; (B) 100 &mu;l/sample of a 3:1 mix of Hyb:PBT; (C) 100 &mu;l/sample of a 1:1 mix of Hyb:PBT; (D) 100 &mu;l/sample of a 1:3 mix of Hyb:PBT; (E) 400 &mu;l/sample of PBT.</li>
 <li>Rinse samples with 100 &mu;l of wash solution A, then perform the wash step with 100 &mu;l/sample of solutions A-D at 56 &deg;C for 15 min each. Then, wash samples 4 times for 5 min with 100 &mu;l/sample of solution E at 56 &deg;C.</li>
 <li>Cool samples to room temperature.</li>
</ol>
4. Probe Detection
Overview: This section describes the antibodies and detection reagents used to detect and visualize the distribution of the RNA probe signal following hybridization. These steps are outlined schematically in Figure 2. Here we only present the standard detection reagents that we use for robust signal amplification, but there exists a variety commercially available reagents that can be used as alternatives, especially when performing double-FISH experiments to co-detect different RNAs simultaneously for protein-RNA double staining. Examples of results obtained with this protocol are displayed in the mRNA expression/localization pattern mosaic in Figure 3.
Materials
<ul>
 <li>HRP-conjugated mouse monoclonal anti-DIG (Jackson ImmunoResearch Laboratories Inc. Cat. No. 200-032-156)</li>
 <li>Biotin-conjugated mouse monoclonal anti-DIG (Jackson ImmunoResearch Laboratories Inc. Cat. No. 200-062-156)</li>
 <li>Streptavidin-HRP conjugate (Molecular Probes, Eugene OR, USA, Cat. No.S991)</li>
 <li>Cy3-tyramide conjugates (Perkin Elmer Life Sciences, Boston, MA, USA, Cat. No. SAT704A)</li>
 <li>DABCO mounting solution: In a 50 ml tube, dilute 1.25 grams of DABCO (1,4-diazabicyclo [2.2.2] octane; Sigma D-2522) in 15 ml of 1X PBS, then add 35 ml of glycerol and mix until fully homogeneous. By premixing with PBS, the DABCO powder is dissolved very quickly. Store mounting solution at -20 &deg;C.</li>
 <li>Glass slides, coverslips</li>
</ul>
Equipment
<ul>
 <li>Nutating mixer</li>
 <li>Multichannel pipettor</li>
 <li>Fluorescence microscope</li>
</ul>
<ol>
 <li>Prepare PBTB solution, containing PBT + 1% non-fat dry milk (usually 50-100 ml is sufficient for all the detection reagent incubation and washing steps).</li>
 <li>Block samples for 10 min at room temp in 100 &mu;l PBTB/sample while rocking on nutator.</li>
 <li>Remove blocking solution from embryos and incubate samples for 1.5-2 hr at room temperature with 100 &mu;l/sample the monoclonal biotin anti-DIG antibody solution (diluted 1/400 from stock into PBTB) with rocking.</li>
 <li>Wash samples 6 times for 5 min each with 100 &mu;l PBTB/sample with rocking.</li>
 <li>Incubate 1.5 hr at room temperature with 75 ml/sample of streptavidin-HRP solution (diluted 1/100 from stock in PBTB, 10 &mu;g/ml final concentration) with rocking.</li>
 <li>Wash samples 6 times for 5 min each 100 &mu;l PBTB/sample with rocking (for the DNA counterstaining, dilute DAPI to 1X working concentration in PBTB and use this solution in the first wash step).</li>
 <li>Rinse embryos with PBT, then wash 2 times for 5 min with PBS.</li>
 <li>Incubate samples for 2 hr at room temperature on the nutator with 50 &mu;l/sample of Cy3-tyramide solution (diluted 1/50 in Tyramide amplification buffer). From this step on, make sure to keep samples in a light shielded receptacle.</li>
 <li>Wash samples 6 times for 5 min each with 100 &mu;l PBS/sample on nutator.</li>
 <li>Remove PBS and add 100-125 &mu;l/sample of mounting solution containing 70% glycerol and 2.5% (DABCO). Store samples at 4 &deg;C. These samples can be stored for several years.</li>
 <li>Using a wide-bore tip, resuspend the embryos by gently pipetting the samples and transfer 25 &mu;l of embryos onto a glass slide, then place a coverslip over the embryos and seal onto the slide with fingernail polish.</li>
 <li>Analyze embryo samples using a fluorescence microscope.</li>
</ol>

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Cytochem. 41, 1725-1733 (1993).</li><li>Tautz, D., Pfeifle, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+non-radioactive+in+situ+hybridization+method+for+the+localization+of+specific+RNAs+in+Drosophila+embryos+reveals+translational+control+of+the+segmentation+gene+hunchback.">A non-radioactive in situ hybridization method for the localization of specific RNAs in Drosophila embryos reveals translational control of the segmentation gene hunchback.</a> Chromosoma. 98, 81-85 (1989).</li><li>Lecuyer, E., Tomancak, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+the+gene+expression+universe.">Mapping the gene expression universe.</a> Curr Opin Genet Dev. 18, 506-512 (2008).</li><li>Tomancak, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Global+analysis+of+patterns+of+gene+expression+during+Drosophila+embryogenesis.">Global analysis of patterns of gene expression during Drosophila embryogenesis.</a> Genome Biol. 8, 2007-208 (2007).</li><li>Thisse, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+and+temporal+expression+of+the+zebrafish+genome+by+large-scale+in+situ+hybridization+screening.">Spatial and temporal expression of the zebrafish genome by large-scale in situ hybridization screening.</a> Methods Cell Biol. 77, 505-519 (2004).</li><li>Quiring, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large-scale+expression+screening+by+automated+whole-mount+in+situ+hybridization.">Large-scale expression screening by automated whole-mount in situ hybridization.</a> Mech. 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No conflicts of interest declared.

For probe synthesis steps, we typically generate run-off antisense RNA probes by in vitro transcription from full-length Drosophila cDNAs amplified from plasmids found in the Drosophila Gene Collection (DGC), a resource detailed at the following website: <a href="http://www.fruitfly.org/DGC/index.html" target="_blank">http://www.fruitfly.org/DGC/index.html</a> 14,15. This approach has been used extensively in large scale ISH studies aimed at mapping gene expression and mRNA localizat...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the Reagent</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>T7, T3 or SP6 RNA Polymerase </td>
 <td>Fermentas Life Sciences </td>
 <td>EP0101,EP0111,EP0131 </td>
 <td>Kits contain reaction buffer. </td>
 </tr>
 <tr>
 <td>DIG RNA Labeling Mix </td>
 <td>Roche Applied Science </td>
 <td>11 277 073 910 </td>
 <td></td>
 </tr>
 <tr>
 <td>RNAguard </td>
 <td>Amersham Biosciences </td>
 <td>27-0816-01 </td>
 <td></td>
 </tr>
 <tr>
 <td>3M sodium acetate </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Cold 100% ethanol. </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Cold 70% ethanol. </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Chlorine bleach solution diluted 1:1 with water. </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Heptane </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Methanol </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>proteinase K </td>
 <td>Sigma Aldrich Oakville, ON, Canada </td>
 <td>Catalog No. P2308 </td>
 <td></td>
 </tr>
 <tr>
 <td>40% formaldehyde solution, freshly prepared </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>PBS-Tween solution (PBT) </td>
 <td></td>
 <td>1xPBS, 0.1% Tween-20 </td>
 <td></td>
 </tr>
 <tr>
 <td>Glycine solution </td>
 <td></td>
 <td>2 mg/ ml glycine in PBT </td>
 <td></td>
 </tr>
 <tr>
 <td>HRP-conjugated mouse monoclonal anti-DIG </td>
 <td>Jackson ImmunoResearch Laboratories Inc </td>
 <td>200-032- 156 </td>
 <td>(1/400 dilution of a 1 mg/ml stock solution in PBTB </td>
 </tr>
 <tr>
 <td>HRP-conjugated sheep monoclonal anti-DIG </td>
 <td>Roche Applied Science, Laval, QC </td>
 <td>1 207 733 </td>
 <td>1/500 dilution of stock solution in PBTB </td>
 </tr>
 <tr>
 <td>Biotin-conjugated mouse monoclonal anti-DIG </td>
 <td>Jackson ImmunoResearch Laboratories Inc., West Grove, PA, USA </td>
 <td>200-062-156 </td>
 <td>(1/400 dilution of a 1 mg/ml stock solution in PBTB </td>
 </tr>
 <tr>
 <td>Streptavidin-HRP conjugate </td>
 <td>Molecular Probes, Eugene OR, USA </td>
 <td>S991 </td>
 <td>(1/100 dilution of a 1 &mu;g/ml stock </td>
 </tr>
 </tbody>
</table>

whole mount rna fluorescent in situ hybridization of drosophila embryos

Assessing the expression pattern of a gene, as well as the subcellular localization properties of its transcribed RNA, are key features for understanding its biological function during development. RNA in situ hybridization (RNA-ISH) is a powerful method used for visualizing RNA distribution properties, be it at the organismal, cellular or subcellular levels 1. RNA-ISH is based on the hybridization of a labeled nucleic acid probe (e.g. antisense RNA, oligonucleotides) complementary to the sequence of an mRNA or a non-coding RNA target of interest 2. As the procedure requires primary sequence information alone to generate sequence-specific probes, it can be universally applied to a broad range of organisms and tissue specimens 3. Indeed, a number of large-scale ISH studies have been implemented to document gene expression and RNA localization dynamics in various model organisms, which has led to the establishment of important community resources 4-11. While a variety of probe labeling and detection strategies have been developed over the years, the combined usage of fluorescently-labeled detection reagents and enzymatic signal amplification steps offer significant enhancements in the sensitivity and resolution of the procedure 12. Here, we describe an optimized fluorescent in situ hybridization method (FISH) employing tyramide signal amplification (TSA) to visualize RNA expression and localization dynamics in staged Drosophila embryos. The procedure is carried out in 96-well PCR plate format, which greatly facilitates the simultaneous processing of large numbers of samples.

When performed successfully, this procedure offers a strikingly enhanced level of detail in the spatio-temporal analysis of gene expression and mRNA localization dynamics during early Drosophila embryogenesis. Indeed, as illustrated in Figure 3A for the classical pair-rule gene runt (run), one can use this protocol to observe gene expression events via the detection of nascent transcript foci in groups of expressing nuclei. In addition, as shown in the embryo mosaic in <s...

Watch this Scientific Journal Video about Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos at JoVE.com

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Here we describe a whole-mount fluorescent in situ hybridization (FISH) protocol for determining the expression and localization properties of RNAs expressed during embryogenesis in the fruit fly, Drosophila melanogaster.

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Assessing  the expression pattern of a gene, as well as the subcellular localization  properties of its transcribed RNA, are key features for ...

whole-mount-rna-fluorescent-situ-hybridization-drosophila

Research

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Biology

18.2K vues.  Institut de Recherches Cliniques de Montréal (IRCM). Nous décrivons ici un ensemble de montage fluorescent In situ Hybridation (FISH) pour la détermination des propriétés d'expression et la localisation des ARN exprimés au cours de l'embryogenèse chez la drosophile, Drosophila melanogaster.

Vidéo: Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. The protocol is a modified version of the standard in situ hybridization using alkaline phosphatase and substrates such as NBT/BCIP and Fast Red 1,2. This protocol utilizes standard digoxygenin and fluorescein labeled probes along with tyramide signal amplification (TSA) 3. The commercially available TSA kits allow flexible experimental design as fluorescence emission from green to far-red can be used in combination with various nuclear stains, such as propidium iodide, or fluorescence immunohistochemistry for proteins. TSA produces a reactive fluorescent substrate that quickly covalently binds to moieties, typically tyrosine residues, in the immediate vicinity of the labeled antisense riboprobe. The resulting staining patterns are high resolution in that subcellular localization of the mRNA can be observed using laser scanning confocal microscopy 3,4. One can observe nascent transcripts at the chromosomal loci, distinguish nuclear and cytoplasmic staining and visualize other patterns such as cortical localization of mRNA. Studies in Drosophila indicate that roughly 70% of mRNAs exhibit specific patterns of subcellular localization that frequently correlate with the function of the encoded protein 5. When combined with computer-aided reconstruction of 3D confocal datasets, our protocol allows the detailed analysis of mRNA distribution with sub-cellular resolution in whole vertebrate embryos.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. We include a propidium iodide nuclear counter-stain to highlight tissue organization.

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology.  Here, we present a high-resolution double ...

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of specific mRNA transcripts within whole mount specimen. This technique (adapted from Albertson and Yelick, 2005) was used in an upper level undergraduate Comparative Vertebrate Biology laboratory classroom at Syracuse University. The first two thirds of the Comparative Vertebrate Biology lab course gave students the opportunity to study the embryology and gross anatomy of several organisms representing various chordate taxa primarily via traditional dissections and the use of models. The final portion of the course involved an innovative approach to teaching anatomy through observation of vertebrate development employing molecular techniques in which WISH was performed on zebrafish embryos. A heterozygous fibroblast growth factor 8 a (fgf8a) mutant line, ace, was used. Due to Mendelian inheritance, ace intercrosses produced wild type, heterozygous, and homozygous ace/fgf8a mutants in a 1:2:1 ratio. RNA probes with known expression patterns in the midline and in developing anatomical structures such as the heart, somites, tailbud, myotome, and brain were used. WISH was performed using zebrafish at the 13 somite and prim-6 stages, with students performing the staining reaction in class. The study of zebrafish embryos at different stages of development gave students the ability to observe how these anatomical structures changed over ontogeny. In addition, some ace/fgf8a mutants displayed improper heart looping, and defects in somite and brain development. The students in this lab observed the normal development of various organ systems using both external anatomy as well as gene expression patterns. They also identified and described embryos displaying improper anatomical development and gene expression (i.e., putative mutants).
For instructors at institutions that do not already own the necessary equipment or where funds for lab and curricular innovation are limited, the financial cost of the reagents and apparatus may be a factor to consider, as will the time and effort required on the part of the instructor regardless of the setting. Nevertheless, we contend that the use of WISH in this type of classroom laboratory setting can provide an important link between developmental genetics and anatomy. As technology advances and the ability to study organismal development at the molecular level becomes easier, cheaper, and increasingly popular, many evolutionary biologists, ecologists, and physiologists are turning to research strategies in the field of molecular biology. Using WISH in a Comparative Vertebrate Biology laboratory classroom is one example of how molecules and anatomy can converge within a single course. This gives upper level college students the opportunity to practice modern biological research techniques, leading to a more diversified education and the promotion of future interdisciplinary scientific research.

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) was used in an upper level undergraduate Comparative Vertebrate Biology course in addition to vertebrate dissections. This gave students the opportunity to study gene expression patterns as well as gross anatomy, linking the study of molecular and organismal biology within one course.

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of ...

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures are lengthy and technically demanding with multiple important steps that collectively contribute to the quality of the final result. This protocol describes in detail several key quality control steps for optimizing probe labeling and performance. Overall, our protocol provides a detailed description of the critical steps necessary to reproducibly obtain high quality results. First, we describe the generation of digoxygenin (DIG) labeled RNA probes via in vitro transcription of DNA templates generated by PCR. We describe three critical quality control assays to determine the amount, integrity and specific activity of the DIG-labeled probes. These steps are important for generating a probe of sufficient sensitivity to detect endogenous mRNAs in a whole mouse embryo. In addition, we describe methods for the fixation and storage of E8.5-E11.5 day old mouse embryos for in situ hybridization. Then, we describe detailed methods for limited proteinase K digestion of the rehydrated embryos followed by the details of the hybridization conditions, post-hybridization washes and RNase treatment to remove non-specific probe hybridization. An AP-conjugated antibody is used to visualize the labeled probe and reveal the expression pattern of the endogenous transcript. Representative results are shown from successful experiments and typical suboptimal experiments.

Whole Mount in Situ Hybridization of E8.5 to E11.5 Mouse Embryos

This whole mount in situ hybridization protocol discusses critical steps that ensure reproducible high quality results for gene expression studies in E8.5-E11.5 day old mouse embryos.

Whole mount in situ hybridization is a very informative approach for defining gene expression patterns in embryos. The in situ hybridization procedures ...

Detection of Viral RNA by Fluorescence in situ Hybridization (FISH)

Viruses that infect cells elicit specific changes to normal cell functions which serve to divert energy and resources for viral replication. Many aspects of host cell function are commandeered by viruses, usually by the expression of viral gene products that recruit host cell proteins and machineries. Moreover, viruses engineer specific membrane organelles or tag on to mobile vesicles and motor proteins to target regions of the cell (during de novo infection, viruses co-opt molecular motor proteins to target the nucleus; later, during virus assembly, they will hijack cellular machineries that will help in the assembly of viruses). Less is understood on how viruses, in particular those with RNA genomes, coordinate the intracellular trafficking of both protein and RNA components and how they achieve assembly of infectious particles at specific loci in the cell. The study of RNA localization began in earlier work. Developing lower eukaryotic embryos and neuronal cells provided important biological information, and also underscored the importance of RNA localization in the programming of gene expression cascades. The study in other organisms and cell systems has yielded similar important information. Viruses are obligate parasites and must utilise their host cells to replicate. Thus, it is critical to understand how RNA viruses direct their RNA genomes from the nucleus, through the nuclear pore, through the cytoplasm and on to one of its final destinations, into progeny virus particles 1.
FISH serves as a useful tool to identify changes in steady-state localization of viral RNA. When combined with immunofluorescence (IF) analysis 22, FISH/IF co-analyses will provide information on the co-localization of proteins with the viral RNA3. This analysis therefore provides a good starting point to test for RNA-protein interactions by other biochemical or biophysical tests 4,5, since co-localization by itself is not enough evidence to be certain of an interaction. In studying viral RNA localization using a method like this, abundant information has been gained on both viral and cellular RNA trafficking events 6. For instance, HIV-1 produces RNA in the nucleus of infected cells but the RNA is only translated in the cytoplasm. When one key viral protein is missing (Rev) 7, FISH of the viral RNA has revealed that the block to viral replication is due to the retention of the HIV-1 genomic RNA in the nucleus 8. 
Here, we present the method for visual analysis of viral genomic RNA in situ. The method makes use of a labelled RNA probe. This probe is designed to be complementary to the viral genomic RNA. During the in vitro synthesis of the antisense RNA probe, the ribonucleotide that is modified with digoxigenin (DIG) is included in an in vitro transcription reaction. Once the probe has hybridized to the target mRNA in cells, subsequent antibody labelling steps (Figure 1) will reveal the localization of the mRNA as well as proteins of interest when performing FISH/IF.

Detection of Viral RNA by Fluorescence in situ Hybridization FISH

A fluorescence in situ hybridization (FISH) method was developed to visually detect viral genomic RNA using fluorescence microscopy. A probe is made with specificity to the viral RNA that can then be identified using a combination of hybridization and immunofluorescence techniques. This technique offers the advantage of identifying the localization of the viral RNA or DNA at steady-state, providing information on the control of intracellular virus trafficking events.

Viruses  that infect cells elicit specific changes to normal cell functions which serve  to divert energy and resources for viral replication. Many ...

Analysis of Single-cell Gene Transcription by RNA Fluorescent In Situ Hybridization (FISH)

Adhesion of Plasmodium falciparum infected erythrocytes (IE) to human endothelial receptors during malaria infections is mediated by expression of PfEMP1 protein variants encoded by the var genes.
The haploid P. falciparum genome harbors approximately 60 different var genes of which only one has been believed to be transcribed per cell at a time during the blood stage of the infection. How such mutually exclusive regulation of var gene transcription is achieved is unclear, as is the identification of individual var genes or sub-groups of var genes associated with different receptors and the consequence of differential binding on the clinical outcome of P. falciparum infections. Recently, the mutually exclusive transcription paradigm has been called into doubt by transcription assays based on individual P. falciparum transcript identification in single infected erythrocytic cells using RNA fluorescent in situ hybridization (FISH) analysis of var gene transcription by the parasite in individual nuclei of P. falciparum IE1.
Here, we present a detailed protocol for carrying out the RNA-FISH methodology for analysis of var gene transcription in single-nuclei of P. falciparum infected human erythrocytes. The method is based on the use of digoxigenin- and biotin- labeled antisense RNA probes using the TSA Plus Fluorescence Palette System2 (Perkin Elmer), microscopic analyses and freshly selected P. falciparum IE. The in situ hybridization method can be used to monitor transcription and regulation of a variety of genes expressed during the different stages of the P. falciparum life cycle and is adaptable to other malaria parasite species and other organisms and cell types.

Analysis of Single-cell Gene Transcription by RNA Fluorescent In Situ Hybridization FISH

Fluorescent in situ hybridization (FISH) to identify mRNA transcripts in individual cells allows analysis of polygenic activity such as the simultaneous transcription of more than one member of the var multigene family in Plasmodium falciparum infected erythrocytes 1. The technique is adaptable and can be used on different types of genes, cells and organisms.

Adhesion of Plasmodium falciparum infected erythrocytes (IE) to human endothelial receptors during malaria infections is mediated by expression of PfEMP1 ...

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The specification of replication timing is thought to be a dynamic process regulated by tissue-specific and developmental cues that are responsive to epigenetic modifications. However, the mechanisms regulating where and when DNA replication initiates along chromosomes remains poorly understood. Homologous chromosomes usually replicate synchronously, however there are notable exceptions to this rule. For example, in female mammalian cells one of the two X chromosomes becomes late replicating through a process known as X inactivation1. Along with this delay in replication timing, estimated to be 2-3 hr, the majority of genes become transcriptionally silenced on one X chromosome. In addition, a discrete cis-acting locus, known as the X inactivation center, regulates this X inactivation process, including the induction of delayed replication timing on the entire inactive X chromosome. In addition, certain chromosome rearrangements found in cancer cells and in cells exposed to ionizing radiation display a significant delay in replication timing of &gt;3 hours that affects the entire chromosome2,3. Recent work from our lab indicates that disruption of discrete cis-acting autosomal loci result in an extremely late replicating phenotype that affects the entire chromosome4. Additional 'chromosome engineering' studies indicate that certain chromosome rearrangements affecting many different chromosomes result in this abnormal replication-timing phenotype, suggesting that all mammalian chromosomes contain discrete cis-acting loci that control proper replication timing of individual chromosomes5.
Here, we present a method for the quantitative analysis of chromosome replication timing combined with fluorescent in situ hybridization. This method allows for a direct comparison of replication timing between homologous chromosomes within the same cell, and was adapted from6. In addition, this method allows for the unambiguous identification of chromosomal rearrangements that correlate with changes in replication timing that affect the entire chromosome. This method has advantages over recently developed high throughput micro-array or sequencing protocols that cannot distinguish between homologous alleles present on rearranged and un-rearranged chromosomes. In addition, because the method described here evaluates single cells, it can detect changes in chromosome replication timing on chromosomal rearrangements that are present in only a fraction of the cells in a population.

Chromosome Replicating Timing Combined with Fluorescent In situ Hybridization

A quantitative method for the analysis of chromosome replication timing is described. The method utilizes BrdU incorporation in combination with fluorescent in situ hybridization (FISH) to assess replication timing of mammalian chromosomes. This technique allows for the direct comparison of rearranged and un-rearranged chromosomes within the same cell.

Mammalian DNA replication initiates at multiple sites along chromosomes at different times during S phase, following a temporal replication program. The ...

RNA In situ Hybridization in Whole Mount Embryos and Cell Histology Adapted for Marine Elasmobranchs

Marine elasmobranchs are valued animal models for biomedical and genomic studies as they are the most primitive vertebrates to have adaptive immunity and have unique mechanisms for osmoregulation 1-3. As the most primitive living jawed-vertebrates with paired appendages, elasmobranchs are an evolutionarily important model, especially for studies in evolution and development. Marine elasmobranchs have also been used to study aquatic toxicology and stress physiology in relationship to climate change 4. Thus, development and adaptation of methodologies is needed to facilitate and expand the use of these primitive vertebrates to multiple biological disciplines. Here I present the successful adaptation of RNA whole mount in situ hybridization and histological techniques to study gene expression and cell histology in elasmobranchs.
Monitoring gene expression is a hallmark tool of developmental biologists, and is widely used to investigate developmental processes 5. RNA whole mount in situ hybridization allows for the visualization and localization of specific gene transcripts in tissues of the developing embryo. The expression pattern of a gene's message can provide insight into what developmental processes and cell fate decisions a gene may control. By comparing the expression pattern of a gene at different developmental stages, insight can be gained into how the role of a gene changes during development.
While whole mount in situ's provides a means to localize gene expression to tissue, histological techniques allow for the identification of differentiated cell types and tissues. Histological stains have varied functions. General stains are used to highlight cell morphology, for example hematoxylin and eosin for general staining of nuclei and cytoplasm, respectively. Other stains can highlight specific cell types. For example, the alcian blue stain reported in this paper is a widely used cationic stain to identify mucosaccharides. Staining of the digestive tract with alcian blue can identify the distribution of goblet cells that produce mucosaccharides. Variations in mucosaccharide constituents on short peptides distinguish goblet cells by function within the digestive tract 6. By using RNA whole mount in situ's and histochemical methods concurrently, cell fate decisions can be linked to gene-specific expression.
Although RNA in situ's and histochemistry are widely used by researchers, their adaptation and use in marine elasmobranchs have met limited and varied success. Here I present protocols developed for elasmobranchs and used on a regular basis in my laboratory. Although further modification of the RNA in situ's hybridization method may be needed to adapt to different species, the protocols described here provide a strong starting point for researchers wanting to adapt the use of marine elasmobranchs to their scientific inquiries.

RNA In situ Hybridization in Whole Mount Embryos and Cell Histology Adapted for Marine Elasmobranchs

By combining methods for RNA whole mount in situ hybridization and histology, gene expression can be linked with cell fate decisions in the developing embryo. These methods have been adapted to marine elasmobranchs and facilitate the use of these animals as model organisms for biomedical, toxicology and comparative studies.

Marine  elasmobranchs are valued animal models for biomedical and genomic studies as  they are the most primitive vertebrates to have adaptive immunity ...

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression both in terms of tissue distribution and developmental stage. Protocols are described for the use of WISH of zebrafish embryos using antisense RNA probes labeled with digoxigenin. Probes are generated by incorporating digoxigenin-linked nucleotides through in vitro transcription of gene templates that have been cloned and linearized. The chorions of embryos harvested at defined developmental stages are removed before incubation with specific probes. Following a washing procedure to remove excess probe, embryos are incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase. By employing a chromogenic substrate for alkaline phosphatase, specific gene expression can be assessed. Depending on the level of gene expression the entire procedure can be completed within 2-3 days.

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

The zebrafish, a small tropical fish, has become a popular model for studying gene function during vertebrate development and disease. The temporal and spatial expression of target genes can be determined by in situ hybridization. Our improved protocol allows for the detection of low abundant transcripts with low non-specific background signal.

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression ...

Analysis of Apoptosis in Zebrafish Embryos by Whole-mount Immunofluorescence to Detect Activated Caspase 3

Whole-mount immunofluorescence to detect activated Caspase 3 (Casp3 assay) is useful to identify cells undergoing either intrinsic or extrinsic apoptosis in zebrafish embryos. The whole-mount analysis provides spatial information in regard to tissue specificity of apoptosing cells, although sectioning and/or colabeling is ultimately required to pinpoint the exact cell types undergoing apoptosis. The whole-mount Casp3 assay is optimized for analysis of fixed embryos between the 4-cell stage and 32 hr-post-fertilization and is useful for a number of applications, including analysis of zebrafish mutants and morphants, overexpression of mutant and wild-type mRNAs, and exposure to chemicals. Compared to acridine orange staining, which can identify apoptotic cells in live embryos in a matter of hours, Casp3 and TUNEL assays take considerably longer to complete (2-4 days). However, because of the dynamic nature of apoptotic cell formation and clearance, analysis of fixed embryos ensures accurate comparison of apoptotic cells across multiple samples at specific time points. We have also found the Casp3 assay to be superior to analysis of apoptotic cells by the whole-mount TUNEL assay in regard to cost and reliability. Overall, the Casp3 assay represents a robust, highly reproducible assay in which to analyze apoptotic cells in early zebrafish embryos.

Certain genetic perturbations or exposure to toxins can disrupt normal developmental processes leading to death of specific cell types. The analysis of activated Caspase 3 by whole-mount immunofluorescence in zebrafish embryos reveals stage- and tissue-specific localization of cells specifically undergoing apoptosis.

Whole-mount immunofluorescence to detect activated Caspase 3 (Casp3 assay) is useful to identify cells undergoing either intrinsic or extrinsic apoptosis ...