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1Institut de Recherches Cliniques de Montréal (IRCM), 2Department of Biochemistry, Université de Montréal
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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.
Keywords: Developmental Biology, Issue 71, Cellular Biology, Molecular Biology, Genetics, Genomics, Drosophila, Embryo, Fluorescent in situ hybridization, FISH, Gene Expression Pattern, RNA Localization, RNA, Tyramide Signal Amplification, TSA, knockout, fruit fly, whole mount, embryogenesis, animal model
Legendre, F., Cody, N., Iampietro, C., Bergalet, J., Lefebvre, F. A., Moquin-Beaudry, G., et al. Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos. J. Vis. Exp. (71), e50057, doi:10.3791/50057 (2013).
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.
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).
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.
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 μl 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.
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.
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 Figure 3B, the method enables the visualization of mRNA localization features in high-resolution.
Figure 1. Outline of RNA probe preparation procedure. (A) Schematic representation of our strategy for synthesizing Dig-labelled RNA probes by in vitro transcription using a linear DNA template flanked with bacteriophage RNA polymerase promoter elements. (B) Picture of standard 1% agarose gel showing examples of in vitro transcribed RNA probes for the histone h2a, run and doc transposon RNAs. We usually perform a side-by-side electrophoresis of both the PCR template and RNA probe. The RNA typically appears as an intense discrete band with higher mobility compared to the template
Figure 2. Schematic representation of the probe detection steps in the FISH method. Following probe hybridization to processed embryos, the Dig-labelled probes are recognized using a biotinylated α-Dig antibody, which is then complexed to HRP-conjugated streptavidin. Tyramide signal amplification (TSA) is then performed resulting in the transformation of the tyramide reagent into free radical form through the activity of HRP. Transiently reactive tyramide free radicals covalently bind to aromatic residues of nearby proteins, preventing their diffusion away from the probe location. The tyramide substrate is complexed to fluorescent dyes, which can be detected by fluorescence microscopy.
Figure 3. Examples of results obtained using this FISH procedure highlighting the diversity of mRNA expression and subcellular localization patterns in Drosophila embryos. (A) FISH analysis of the pair-rule gene run at distinct stages of embryogenesis reveals how the initial broad expression in the mid-portion of the embryo at embryonic stage 4 is refined into a segmented stripe pattern in later stages. (B) Mosaic depiction of FISH results showing different varieties of mRNA localization patterns in stage 4-7 embryos. FISH was performed using Dig-labeled antisense probes that were hybridized and detected by sequential incubations with the biotinylated anti-Dig antibody, streptavidin-HRP, and Cy3 tyramide. RNA = blue, DNA = red. Scale bar in (A) = 25 μM, while that in (B) = 50 μM.
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: http://www.fruitfly.org/DGC/index.html 14,15. This approach has been used extensively in large scale ISH studies aimed at mapping gene expression and mRNA localization patterns in fly embryos 9,16. DGC plasmids can be ordered from the Drosophila Genomic Resource Center (https://dgrc.cgb.indiana.edu/vectors/). The flexibility in starting materials and designing custom primers with T7, SP6, or T3 overhangs allows one to easily generate probes to detect specific mRNA isoforms (e.g. alternatively spliced variants) or non-coding RNAs that are not represented in the standard cDNA collections. When considering designing isoform-specific probes, we have found that templates ranging from 250-1,000 bases can yield excellent FISH results.
For FISH in fly embryos, we do not fragment our probes by carbonate treatment; rather we use full-length run-off transcripts, which can range in size between 500-5,000 bases. When performing FISH on other tissues that are less permeable to large probes, fragmentation may indeed favor probe penetration. However, we prefer the option of preparing probes using smaller PCR-amplified DNA templates, or pooling multiple individually-synthesized small probes together, rather than performing chemical fragmentation of large probes which may give variable results and reduce overall probe quality.
As described in the embryo collection steps, we usually add yeast paste to the embryo collection plates. Yeast paste will greatly increase the number of eggs laid by the flies, as egg production depends on the nutritional environment17. In addition, Drosophila flies will frequently lay their eggs directly into the yeast paste. These embryos can easily be collected by dissolving the yeast paste in water using a small paint brush and passing the mixture through a collection basket. For the dechorionation step, a 3% bleach solution, prepared by diluting commercial bleach (typically 6% concentration) 1:1 with water, is used in most protocols 13,18,19. In our experience, we have found that incubating embryos with a 3% bleach solution for 90 sec offers efficient dechorionation. These parameters may need to be adjusted depending on the commercial bleach solution that is available, but one should take care not to overexpose the embryos to bleach since this can damage the samples. To monitor the dechorionation procedure more closely, it is possible to look at the embryos under a microscope to make sure the dechorionation reaction is complete, i.e. the embryos lose their dorsal appendages when they are dechorionated. Finally, to ensure proper embryo fixation, we use freshly prepared formaldehyde solutions, as old solutions tend to precipitate. For further reference, this procedure has been described in previous methods articles 13,18,19.
For steps pertaining to permeabilization of embryonic tissues with proteinase K, or detection reagents such as the antibodies and Cy3-Tyramide, we have found optimal results using the dilutions described in this protocol. However, due to variations in lab environments and reagent stocks, we recommend that each laboratory empirically test their own reagents to determine the best working concentrations for their specific needs. In addition, we have found the optimal reagent concentrations may vary depending on the tissues analyzed.
To ensure that the FISH procedure is giving consistently reproducible results, we recommend always adding several positive controls to the assay, which could include probes for transcripts with well-defined expression/localization patterns (e.g. run). Transcripts with striking mRNA localization patterns during early Drosophila embryogenesis can be found on the Fly-FISH web site (http://fly-fish.ccbr.utoronto.ca/). Conversely, to control for background signal detection, as mentioned above, one should also include 'no-probe' and/or sense RNA probe samples.
In addition to the Cy3 tyramide conjugate described in this protocol, there are a variety of other commercially available fluorochrome-conjugated tyramide reagents from Perkin Elmer Life Sciences or Molecular Probes (Invitrogen, Canada, Inc), which can provide additional flexibility for multicolor imaging experiments. While this method describes our basic procedure for single RNA FISH, variations of this method can be implemented for more complex experiments, such as double-RNA FISH or RNA-protein co-detection. For details pertaining to these procedures, we recommend following the procedures described in existing FISH methods papers 18-20.
No conflicts of interest declared.
Work conducted in the Lécuyer 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 Santé du Québec (FRSQ). Fabio Alexis Lefebvre and Gaël Moquin-Beaudry are supported by NSERC undergraduate research studentships, while Carole Iampietro is supported by the Angelo Pizzagalli postdoctoral fellowship.
|T7, T3 or SP6 RNA Polymerase||Fermentas Life Sciences||EP0101,EP0111,EP0131||Kits contain reaction buffer.|
|DIG RNA Labeling Mix||Roche Applied Science||11 277 073 910|
|3M sodium acetate|
|Cold 100% ethanol.|
|Cold 70% ethanol.|
|Chlorine bleach solution diluted 1:1 with water.|
|proteinase K||Sigma Aldrich Oakville, ON, Canada||Catalog No. P2308|
|40% formaldehyde solution, freshly prepared|
|PBS-Tween solution (PBT)||1xPBS, 0.1% Tween-20|
|Glycine solution||2 mg/ ml glycine in PBT|
|HRP-conjugated mouse monoclonal anti-DIG||Jackson ImmunoResearch Laboratories Inc||200-032- 156||(1/400 dilution of a 1 mg/ml stock solution in PBTB|
|HRP-conjugated sheep monoclonal anti-DIG||Roche Applied Science, Laval, QC||1 207 733||1/500 dilution of stock solution in PBTB|
|Biotin-conjugated mouse monoclonal anti-DIG||Jackson ImmunoResearch Laboratories Inc., West Grove, PA, USA||200-062-156||(1/400 dilution of a 1 mg/ml stock solution in PBTB|
|Streptavidin-HRP conjugate||Molecular Probes, Eugene OR, USA||S991||(1/100 dilution of a 1 μg/ml stock|
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