The authors would like to acknowledge the CIHR for fellowship support of Steve Deimling and the Department of Paediatrics, University of Western Ontario for support of Steve Deimling, Rami Halabi and Stephanie Grover. This work was supported by the NSERC grant R2654A11 and an NSERC Discovery Accelerator Supplement

1. Embryo Preparation
<ol>
	<li>If not done routinely as part of embryo culture, de-jelly the embryos using 2.5% cysteine, pH 8.0 prior to fixation 8. Although it is not absolutely necessary, it is useful to then manually remove the fertilization envelope prior to fixation using fine forceps.
	<ol>
		<li>Use glass Pasteur pipettes to transfer the embryos. The pipette is not wide enough to transfer embryos so use a diamond pen to cut the glass pipette at a point wide enough to pick up an embryo. Eliminate the sharp edges of the pipettes after cutting by quickly passing the cut tip through the flame of a Bunsen burner to melt the sharp edges. 
		NOTE: Care needs to be taken, as the glass can still be hot enough to cause burns even though it appears to have cooled by visual inspection.</li>
	</ol>
	</li>
	<li>Perform the embryo fixation in stages. First, prepare glass vials for use in embryo fixation. Use these vials for all steps including final storage. Use vials that are clear with a good Teflon seal in the lid, allowing for monitoring the embryos during all steps of the process. Label the vials with appropriate experimental information using permanent marker and then cover the label with clear tape, as even permanent marker will be lost over the course of the procedure due to the use of alcohols and other solvents.
	<ol>
		<li>Use Mempfa to fix the embryos. Make a stock of 8% paraformaldehyde in batches of 50-100 ml at a time. Use approximately 75% of the required H2O and heat to 50-60 &#176;C, which is needed to get the paraformaldehyde into solution. 
		NOTE: This solution is slightly different than the Memfa used in older published protocol versions in that it utilizes paraformaldehyde rather than formaldehyde.</li>
		<li>Add 2-3 drops of 10N NaOH or until pH is approximately 7.5 (use pH paper to check pH). Paraformaldehyde is very toxic, therefore generate the stock solution in a fume hood. Once the paraformaldehyde is in solution, filter the solution through Whatman paper into a fresh container and add H2O to the final volume. If required, store the paraformaldehyde solution at 4 &#176;C for 1-2 weeks.</li>
		<li>Assemble the remaining components of the Mempfa fixation solution (Table 1). Ensure that the final working concentration of paraformaldehyde is 4%. Store all components of the fixation solution at 4 &#176;C as stocks.</li>
		<li>Using a cut glass pipette, fix the embryos by adding the embryos to the labeled glass vials that have been filled with the approximately 3-4 ml of Mempfa solution (Table 1). Avoid fixing more than 20-30 embryos per vial. Add the embryos with a minimum of liquid transfer from the embryo medium. Fix embryos in Mempfa solution for 2 hr at room temperature or overnight at 4 &#176;C.</li>
		<li>For late endoderm structures perform the in situs on manually isolated gut and endoderm derivatives 9,10, which allows for good penetration of the probe and also avoids cavity staining.</li>
		<li>Store a solution of 100% methanol at -20 &#176;C. After the paraformaldehyde fixation, replace the Mempfa solution with approximately 4 ml of the -20 &#176;C, 100% methanol for storage of the embryos.</li>
		<li>Swirl the vials after addition of the methanol to prevent the embryos from sticking to the glass or other embryos. Also, make sure the vials are tightly sealed because the methanol can evaporate in the freezer over time if loose. 
		NOTE: Embryos can be stored for at least a year, and likely longer, in the methanol prior to staining with no loss of in situ hybridization quality.</li>
	</ol>
	</li>
</ol>
2. Probe Preparation
<ol>
	<li>Use 1-2 &#181;g of template DNA to make the digoxigenin-labelled probes. Cut plasmid containing the appropriate DNA sequence at the 5&#8217; end of the gene of interest with a restriction enzyme suitable for that vector to generate antisense probes. 
	NOTE: The plasmid should have an appropriate RNA polymerase binding site (e.g. T7, T3, or SP6).</li>
	<li>Allow the DNA, water, NTP mix and polymerase buffer to warm to room temperature before assembling the probe synthesis reaction. Add the components of the RNA synthesis reaction to a 1.5 ml microcentrifuge tube in the order as listed in Table 2. Adjust the volume of water added to bring the total volume of the probe synthesis reaction to 20 &#181;l. Assemble the reaction at room temperature because components of the polymerase buffer can precipitate the template DNA when in high concentrations and cold.</li>
	<li>Incubate the transcription reaction for 2 hr at 37 &#176;C. 
	NOTE: Incubations of slightly longer than two hours leads to no adverse effects, but also show little increased yield. If incubated for one hour, the yield will be reduced but still sufficient to make a good probe.
	<ol>
		<li>While waiting for the transcription reaction to finish, make an agarose gel that will be used to test the quality of the probe. Melt 1 &#181;g of agarose in 100 ml of 1x TAE buffer (see Table 1) by heating the solution to the boiling point. Remove the solution from heat when the agarose powder has completely dissolved.</li>
		<li>Add 2 &#181;l ethidium bromide stock solution (10 mg/ml) to about 100 ml of agarose gel when the agarose has cooled to about 60 &#176;C to allow visualization of RNA under ultraviolet (UV) light. 
		NOTE: This agarose gel solution can be kept in a 60 &#176;C incubator so that future gels can be poured without repeated melting. Take care in handling ethidium bromide due to potential toxicity.</li>
	</ol>
	</li>
	<li>Add 1 &#181;l DNAseI (RNAse free grade) to the transcription reaction after the 2 hr incubation and incubate for a further 10 min at 37 &#176;C to eliminate template DNA.</li>
	<li>Remove 1 &#181;l of the reaction mix to check on the 1% agarose-TAE gel and to the rest (20 &#181;l) add 80 &#181;l of 1% SDS in TE buffer (10 mM Tris pH 8.0, 1 mM EDTA), 10 &#181;l of 5M NH4 Acetate, and 220 &#181;l of cold ethanol. Vortex the mix vigorously and set aside on ice until the results of the RNA quality check are known.
	<ol>
		<li>Run the 1 &#181;l of RNA removed before precipitation on the 1% agarose-TAE gel. To ease loading on the gel, add 4-5 &#181;l of water and 1 &#181;l of standard loading dye to the RNA. View the RNA on the gel using a UV light transilluminator in order to check the quality of the probe (see discussion).</li>
	</ol>
	</li>
	<li>Precipitate the remaining RNA that was set aside on ice in step 2.5 by spinning in a microcentrifuge at full speed for 10 to 15 min. Draw off the supernatant with a drawn out glass pipette and allow to dry briefly.
	<ol>
		<li>Resuspend the probe with 1 ml of RNA hybridization buffer (Table 1) in the eppendorf tube.&#160; Vortex and briefly heat the tube to 37 &#176;C and vortex again.&#160; Transfer the probe solution to a 15 ml screw cap polystyrene tube and fill to 7-10 ml with RNA hybridization buffer.&#160; Note: The probe can be diluted further (a 10 fold further dilution can still work) often resulting in low background but the staining reactions will also take longer.</li>
	</ol>
	</li>
</ol>
3. In situ Hybridization
<ol>
	<li>Take the embryos that were stored in -20 &#176;C methanol and allow to warm to room temperature. Perform the entire procedure in the glass vials. If different embryos from one group are going to be looked at by different probes, keep them in a single vial until just before the probes are added to reduce variability and labor on the first day.
	<ol>
		<li>Rehydrate the embryos through a methanol series as outlined in Table 3 (see Table 1 for wash recipes) in preparation for addition of the probe. Gently swirl the embryos after each change to ensure they are not sticking to the sides or each other, and rock the embryos by mounting the vials on a nutator. When transferring the liquids, check the inside of the caps to ensure that embryos have not been trapped there.</li>
		<li>Once in the probe solution, hybridize the embryos overnight as described in Table 3.</li>
	</ol>
	</li>
	<li>Remove the probe solution. Save the used probe by storing in a 15 ml screw cap polystyrene tube, marked with date and number of times the probe has been used, at -20 &#176;C. 
	NOTE: The same probe can be reused many times for subsequent in situ hybridizations until the colorimetric reactions begin to take an abnormally long time to reach desired intensity.
	<ol>
		<li>Once the probe is removed, prepare the embryos for antibody staining against the probe through the series of washes outlined in Table 4. Change the temperature as required (Table 4) by moving the nutator, with vials of embryos attached, directly into hybridization ovens that are set to the appropriate temperature.</li>
		<li>Make up the appropriate volume of the MAB+HTSS+BR+anti-Dig antibody (Table 4) at the time that the embryos are being incubated in the MAB+HTSS+BR blocking solution so that the antibody is blocked prior to adding to the embryos. Make the blocking solutions fresh on the day of use.</li>
	</ol>
	</li>
	<li>Remove the antibody solution and begin washes as outlined in Table 5 following overnight incubation with antibody. Perform at least twelve 30 min washes in order to prepare for staining with the alkaline phosphatase substrate and reduce the background as much as possible. Use either MAB buffer or TBT solution (Table 1) for the washing steps. 
	NOTE: TBT solution is TTw that has 2 mg/ml of BSA added.
	<ol>
		<li>Replace the last wash solution with the BM Purple alkaline phosphatase substrate. Perform the staining reaction at either room temperature or 37 &#176;C. 
		NOTE: Staining is more rapid at 37 &#176;C but if left overnight, unacceptable background staining can often be a result. The staining reactions will often require overnight staining and room temperature is safest for this.</li>
		<li>If further staining is required and the BM purple solution is taking on a blue color, replace the staining solution with fresh BM Purple and put the tubes into 37 &#176;C. If the target mRNA is very abundant, put the embryos in the staining solution at 4 &#176;C overnight and then move the embryos to room temperature or 37 &#176;C to allow better monitoring of the staining reaction.</li>
		<li>Monitor new reactions carefully, as the time to final result varies considerably between different target RNAs. For consistency, stop the staining reaction (see 3.4) when there are no new sites of expression arising with longer incubation. Importantly, if in situ hybridization is being used as an assay for experiments looking at different treatment groups, use the same time for color reactions between treatment and control embryos.</li>
	</ol>
	</li>
	<li>Stop the staining reaction and prepare the embryos for storage and image recording by changing the liquids as outlined in Table 6. Do not rock the embryos at this stage because the removal of stain can be visualized as purple coloring of the methanol around the embryos. 
	NOTE: Often embryos have a light blue staining from the staining solution and the cold methanol can at least partially remove that, although this is not sufficient to remove heavy background or cavity staining. Cold methanol refers to methanol that is maintained in a -20 &#176;C freezer.
	<ol>
		<li>Rehydrate the embryos and fix the stain with Mempfa (Table 6). Once fixed, remove the Mempfa and wash the embryos with 25% methanol.</li>
		<li>Remove the 25% methanol and add the bleaching solution if removal of endogenous pigment is required. Take care in handling the bleaching solution as it can cause burns. Observe the bleaching closely as it happens relatively quickly and the degree of bleaching can be varied for different effects (Figure 2).</li>
		<li>Dehydrate embryos through a methanol series to 100% methanol for long term storage following bleaching, or transfer to PBS for short term storage and subsequent imaging (see Table 6).&#160;</li>
	</ol>
	</li>
</ol>
4. Imaging Embryos
<ol>
	<li>Once an embryo has completed the staining process, image the embryo so that the information on where the gene of interest is expressed is captured for a wider audience. Use 1% agarose as a background to view uncleared embryos. 
	NOTE: The agarose gives a blue/grey background that contrasts well with the embryo and the blue color of the staining reaction. It also helps diffuse distracting shadows and reflections that divert attention from the embryo.
	<ol>
		<li>Add agarose to water and then bring to a boil until the agarose is in solution and then let cool to 50 before pouring into the petri dish. As with the TAE gel solution, store the agarose solution in 55 &#176;C incubator for multiple uses. Pour the agarose to a depth of about 2 mm into the petri dish. If required, adjust the depth of agarose to yield a slightly differently shade of background.</li>
		<li>Following rehydration from storage in methanol to an aqueous solution (PBS or TTw), using a methanol series, place the embryos in a petri dish with the agarose base (see Table 6). Keep the solution clean and if required, use simple filtering to eliminate small particulate matter that can disrupt the clean background of good images.</li>
		<li>To image the embryos from alternate views, such as from the ventral side, cut thin channels in the agarose to fit the embryo using fine forceps and place the embryos in those channels for orientation (Figure 3). Take care in manipulating the embryos as they are easily damaged. 
		NOTE: Most stages have a characteristic position that they assume when placed in solution. For example, blastula-stage embryos tend to sit animal side up. Once the embryos begin to elongate, they lay on their side.</li>
	</ol>
	</li>
	<li>Use the fiber optic light source to illuminate the embryo from a shallow angle, creating shadows on the embryo that provide depth to the image and help discern surface structures. Because bleaching can eliminate pigment that provides useful landmarks, use shadowing for strongly bleached embryos.</li>
	<li>Clear the embryos to image staining that is deep within the embryo, such as in the notochord, lung, or regions of the brain, (Figure 4). To accomplish this, put the embryos through a methanol series until in 100% methanol.
	<ol>
		<li>After complete immersion in methanol, transfer the embryos to a solution of one part benzyl alcohol and two parts benzyl benzoate (BABB). The embryos will initially float on the surface but as the methanol mixes with the BABB, they will sink into the BABB. Perform every step dealing with BABB in glass vials or dishes; BABB will melt any plastic or paint.</li>
		<li>Once cleared, view the embryos with transmitted light coming from below the embryo. Adjust the intensity of the light as well as the angle from below to improve contrast and provide the best color.</li>
		<li>When viewing cleared embryos raise the glass petri dish off of the base. Do this by simply using two other petri dish lids to raise it so that the region of the dish with embryos is elevated. 
		NOTE: This has the advantage of taking the base out of focus and eliminating distracting effects from imperfections or stains on the microscope base that can interfere with the image.</li>
	</ol>
	</li>
</ol>
5. Double In situ Hybridization
<ol>
	<li>In order to simultaneously view the expression pattern of two different genes in a single embryo, synthesize two probes, one for each of the different genes. Synthesize one probe using DIG-11-UTP as a label as described above. Dilute the product of the transcription reaction in RNA hybridization buffer to yield a 3x more concentrated probe than used for single in situ hybridizations.
	<ol>
		<li>Synthesize the other probe of interest using the same protocol as for DIG labelled probed except that fluorescein-12-UTP must be substituted for DIG-11-UTP. Dilute the product of the transcription reaction in RNA hybridization buffer to yield a 3x more concentrated probe than used for single in situ hybridizations.</li>
		<li>Mix the two concentrated probes in a 1:1 ratio. For best results, use the fluorescein-labelled probe for the gene that shows the strongest expression in the single in situ hybridization protocol.</li>
	</ol>
	</li>
	<li>Use the same in situ hybridization protocol described for single in situ hybridizations, except use the double probe (probe containing the 1.5x concentrated mixture of digoxigenin-labelled and fluorescein-labelled probes) at the end of the first day in place of a single in situ hybridization probe.
	<ol>
		<li>Follow the single hybridization protocol on the second day of the double in situ hybridization protocol, except use anti-fluorescein-AP Fab fragments at a 1:4,000 dilution in place of anti-DIG-AP Fab fragments. Wash out the excess antibody from the embryo as in the single in situ protocol and carry out the first color reaction using the BM-Purple AP substrate.</li>
	</ol>
	</li>
	<li>Following the first color reaction, inactivate the flourescein antibody in 0.1 M glycine pH 2.0 for 40 min followed by five ten min washes in MAB. Block the embryos in MAB+HTSS+BR for 90 min. Add the anti-DIG antibody at a 1:2,000 dilution in MAB+HTSS+BR and incubate at 4 &#176;C overnight.
	<ol>
		<li>The following day, wash the embryos thoroughly in MAB (12 washes of 30 min) to remove the excess antibody.</li>
		<li>Wash the embryos for 10 min in AP Buffer (Table 1) and then stain with BCIP (0.5mg/ml in AP Buffer). 
		NOTE: The in situ combination should give a dark blue-purple stain for the first color reaction and a light blue color reaction for the second (Figure 5).</li>
		<li>Stop the final color reaction by removing the AP buffer and rinse three times with MAB. Fix the embryos with Mempfa for 10 min. Wash the embryos with 5 quick washes in MAB or TBT.</li>
		<li>With this color combination, the use of methanol in the post staining treatments is no longer possible, as it will eliminate the BCIP alone color. Store the embryos after staining and fixation in PBS with 0.02% sodium azide. Staining intensity can be weak with double in situs. If this is a problem, reduce the washes to four, each of 2 hr duration.</li>
	</ol>
	</li>
</ol>

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Authors have no competing financial interests to disclose.

The ability to use in situ hybridization to visualize the expression pattern of specific genes remains the most commonly used method to identify specific organs or cell types in the Xenopus embryo. This is because of several advantages offered by this technique. The expression of a gene can identify specific structures well before any histological sign of differentiation such as the case for nkx2.5 expression in the heart progenitors prior to any clear demarcation of those cells 18. ...

The expression pattern of a specific gene is an important piece of information in determining the potential role for that gene in the development of a specific organ or cell type. Simply put, if it is not expressed at the right time and place it is unlikely to play a key role. In Xenopus, as in most early embryos, the most commonly used assay for detecting the expression of a gene is whole mount in situ hybridization using labeled antisense RNA probes. The use of antibody staining to assess expression of a gene in Xenopus is becoming more common as researchers discover antibodies, usually raised against mammalian proteins, that cross react to the Xenopus homologue or generate their own 1-3. However, the vast majority of studies on Xenopus organogenesis still utilize antisense RNA probes. When antibodies are used, each individual antibody often requires optimization for the primary antibody concentration or fixation protocols. In contrast, the protocol for in situ hybridizations is essentially invariant for different probes. The basic concept is relatively simple and an excellent standard protocol has been well established 4. Our protocol is a streamlined version of the original protocol 4 that still provides excellent detection of gene expression patterns in the early embryo. The embryos are fixed and then prepared for hybridization by changing solutions and temperatures such that it allows for high stringency binding of the labeled antisense RNA probe to its target mRNA. The unbound probe is washed away and the embryos are then prepared for binding of an antibody against the label on the RNA probes. Excess antibody is then washed away and an enzymatic color reaction is used to localize where the RNA probe is bound in the embryo. There are now a number of Xenopus transgenic lines that drive expression of fluorescent proteins in specific tissues and these are available at the Xenopus stock centers such as the National Xenopus Resource in Woods Hole. While very useful for many experiments that require examining organogenesis in living embryos, this option requires separate housing for the transgenic lines. 
In situ hybridization can clearly delineate where specific organs or cell types will form in the early embryo (Figure 1). The technique is remarkably sensitive given that one can detect gene expression in a small number of cells in a single embryo 5. However, in situ hybridization using the intensity of colorimetric staining is not considered quantifiable because the color reaction is not a linear one. Despite difficulty in quantifying staining intensity, changes in expression are often quite noticeable; particularly when the in situ hybridization shows quantifiable increases or decreases in the size of expression domains 6,7. 
The clear advantages of whole mount in situ hybridization make it a critical assay in the study of early development. However, it is a time consuming one that requires many steps over several days. This protocol is a simplified version of the standard protocol that eliminates several steps without reducing the quality of the in situ result. The simplification also eliminates sources of variability, making trouble shooting easier if an in situ hybridization is not optimal. Specifically, we have eliminated the use of proteinase K and RNAse treatments of the embryo, two steps that can depend on reagent quality and can also reduce signal intensity if overdone. The protocol also provides some degree of cost saving due to eliminating the use of several reagents. Finally, this protocol also provides some simple guidelines for improved capturing of images of in situ hybridization results. Although this protocol is optimized for work in Xenopus embryos, it is likely that at least some of the simplifications will be applicable to in situ hybridization work in other embryo systems.

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understanding early organogenesis using a simplified in situ hybridization protocol in xenopus

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development. The Xenopuslaevis embryo is very useful for studying organogenesis because their large size makes them very suitable for identifying organs at the earliest steps in organogenesis. At this time, the primary method used for identifying a specific organ or primordium is whole mount in situ hybridization with labeled antisense RNA probes specific to a gene that is expressed in the organ of interest. In addition, it is relatively easy to manipulate genes or signaling pathways in Xenopus and in situ hybridization allows one to then assay for changes in the presence or morphology of a target organ. Whole mount in situ hybridization is a multi-day protocol with many steps involved. Here we provide a simplified protocol with reduced numbers of steps and reagents used that works well for routine assays. In situ hybridization robots have greatly facilitated the process and we detail how and when we utilize that technology in the process. Once an in situ hybridization is complete, capturing the best image of the result can be frustrating. We provide advice on how to optimize imaging of in situ hybridization results. Although the protocol describes assessing organogenesis in Xenopus laevis, the same basic protocol can almost certainly be adapted to Xenopus tropicalis and other model systems.

The use of tissue specific probes can provide outstanding information in regards to the state of development for specific organs. In the following examples, the stage of the embryo is based on the Nieuwkoop and Faber staging table 11. If one uses probes form genes expressed after differentiation, cardiac troponin I at stage 28-30, for example (Figure 1C), the presence or size of a differentiated organ can be assessed at any stage post differentiation. Years ago, embryologists were abl...

Watch this Scientific Journal Video about Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus at JoVE.com

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

The Xenopus laevis embryo continues to be exceptionally useful in the study of early development due to its large size and ease of manipulation. A simplified protocol for whole mount in situ hybridization protocol is provided that can be used in the identification of specific organs in this model system.

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

Organogenesis is the study of how organs are specified and then acquire their specific shape and functions during development.  The Xenopuslaevis embryo ...

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Developmental Biology

39.0K Views.  Hospital for Sick Children. The Xenopus laevis embryo continues to be exceptionally useful in the study of early development due to its large size and ease of manipulation. A simplified protocol for whole mount in situ hybridization protocol is provided that can be used in the identification of specific organs in this model system.

Video: Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, cells from the roof of the blastocoel are pluripotent. These cells can be isolated, and programmed to generate various tissues through manipulation of genes expression or induction by morphogens. In this manuscript protocols are described for the use of Xenopus laevis blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development. These protocols allow the investigation of fate acquisition, cell migration behaviors, and cell autonomous and non-autonomous properties. The blastocoel roof explants can be cultured in a serum-free defined medium and grafted into host embryos. This transplantation into an embryo allows the investigation of the long-term lineage commitment, the inductive properties, and the behavior of transplanted cells in vivo. These assays can be exploited to investigate molecular mechanisms, cellular processes and gene regulatory networks underlying neural development. In the context of regenerative medicine, these assays provide a means to generate neural-derived cell types in vitro that could be used in drug screening.

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

In Xenopus embryos, cells from the roof of the blastocoel are pluripotent and can be programmed to generate various tissues. Here, we describe protocols to use amphibian blastocoel roof explants as an assay system to investigate key in vivo and in vitro features of early neural development.

Understanding the genetic programs underlying neural development is an important goal of developmental and stem cell biology. In the amphibian blastula, ...

Functional Cloning Using a Xenopus Oocyte Expression System

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in abundance, may not be secreted but tethered to the signaling cell(s), or may require the presence of binding partners or upstream regulatory molecules. Thus in a search for gene products capable of eliciting an early lens-inductive response in competent ectoderm, we utilized an expression cloning system that would allow identification of paracrine or juxtacrine factors as well as transcriptional or other regulatory proteins. Pools of mRNA were injected into Xenopus oocytes, and responding tissue placed directly on the oocytes and co-cultured. Following functional cloning of ldb1 from a neural plate stage cDNA library based on its ability to elicit the expression of the early lens placode marker foxe3 in lens-competent animal cap ectoderm, we characterized the mRNA expression pattern, and assayed developmental progression following overexpression or knockdown of ldb1. This system is suitable in a very wide variety of contexts where identification of an inducer or its upstream regulatory molecules is sought using a functional response in competent tissue.

Functional Cloning Using a Xenopus Oocyte Expression System

We describe a Xenopus oocyte and animal cap system for the expression cloning of genes capable of inducing a response in competent ectoderm, and discuss techniques for the subsequent analysis of such genes. This system is useful in the functional identification of a wide range of gene products.

Identification of genes responsible for embryonic induction poses a number of challenges; to name a few, secreted molecules of interest may be low in ...

Technique to Target Microinjection to the Developing Xenopus Kidney

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus embryos are large, develop externally, and can be easily manipulated by microinjection or surgical procedures. In addition, fate maps have been established for early Xenopus embryos. Targeted microinjection into the individual blastomere that will eventually give rise to an organ or tissue of interest can be used to selectively overexpress or knock down gene expression within this restricted region, decreasing secondary effects in the rest of the developing embryo. In this protocol, we describe how to utilize established Xenopus fate maps to target the developing Xenopus kidney (the pronephros), through microinjection into specific blastomere of 4- and 8-cell embryos. Injection of lineage tracers allows verification of the specific targeting of the injection. After embryos have developed to stage 38 - 40, whole-mount immunostaining is used to visualize pronephric development, and the contribution by targeted cells to the pronephros can be assessed. The same technique can be adapted to target other tissue types in addition to the pronephros.

Technique to Target Microinjection to the Developing Xenopus Kidney

Here, we present a protocol to use fate maps and lineage tracers to target injections into individual blastomeres that give rise to the kidney of Xenopus laevis embryos.

The embryonic kidney of Xenopus&#160;laevis (frog), the pronephros, consists of a single nephron, and can be used as a model for kidney disease. Xenopus ...

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are expressed in spatial relation to each other in situ. Multi-target chromogenic whole-mount in situ hybridization (MC-WISH) greatly facilitates the instant comparison of gene expression patterns, as it allows distinctive visualization of different mRNA species in contrasting colors in the same sample specimen. This provides the possibility to relate gene expression domains topographically to each other with high accuracy and to define unique and overlapping expression sites. In the presented protocol, we describe a MC-WISH procedure for comparing mRNA expression patterns of different genes in Drosophila embryos. Up to three RNA probes, each specific for another gene and labeled by a different hapten, are simultaneously hybridized to the embryo samples and subsequently detected by alkaline phosphatase-based colorimetric immunohistochemistry. The described procedure is detailed here for Drosophila, but works equally well with zebrafish embryos.

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

We describe a multi-target chromogenic whole-mount in situ hybridization (MC-WISH) procedure in intact Drosophila embryos allowing simultaneous and specific detection of three different mRNA distribution patterns by contrasting color precipitates.

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are ...

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and optical transparency. In particular, the zebrafish embryo has become an important organism to study vertebrate kidney organogenesis as well as multiciliated cell (MCC) development. To visualize MCCs in the embryonic zebrafish kidney, we have developed a combined protocol of whole-mount fluorescent in situ hybridization (FISH) and whole mount immunofluorescence (IF) that enables high resolution imaging. This manuscript describes our technique for co-localizing RNA transcripts and protein as a tool to better understand the regulation of developmental programs through the expression of various lineage factors.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and ...

Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk of conceiving an aneuploid fetus increases dramatically and this phenomenon is known as the maternal age effect. One requirement for accurate chromosome segregation during the meiotic divisions is maintenance of sister chromatid cohesion during the extended prophase period that oocytes experience. Cytological evidence in both humans and model organisms suggests that meiotic cohesion deteriorates during the aging process. In addition, segregation errors in human oocytes are most prevalent during meiosis I, consistent with premature loss of arm cohesion. The use of model organisms is critical for unraveling the mechanisms that underlie age-dependent loss of cohesion. Drosophila melanogaster offers several advantages for studying the regulation of meiotic cohesion in oocytes. However, until recently, only genetic tests were available to assay for loss of arm cohesion in oocytes of different genotypes or under different experimental conditions. Here, a detailed protocol is provided for using fluorescence in situ hybridization (FISH) to directly visualize defects in arm cohesion in prometaphase I and metaphase I arrested Drosophila oocytes. By generating a FISH probe that hybridizes to the distal arm of the X chromosome and collecting confocal Z stacks, a researcher can visualize the number of individual FISH signals in three dimensions and determine whether sister chromatid arms are separated. The procedure outlined makes it possible to quantify arm cohesion defects in hundreds of Drosophila oocytes. As such, this method provides an important tool for investigating the mechanisms that contribute to cohesion maintenance as well as the factors that lead to its demise during the aging process.

Using Fluorescence In Situ Hybridization FISH to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

This manuscript presents a detailed method for generating X-chromosome arm probes and performing fluorescence in situ hybridization (FISH) to examine the state of sister chromatid cohesion in prometaphase and metaphase I arrested Drosophila oocytes. This protocol is suitable for determining whether meiotic arm cohesion is intact or disrupted in different genotypes.

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk ...

Two-step Approach to Explore Early- and Late-stages of Organ Formation in the Avian Model: The Thymus and Parathyroid Glands Organogenesis Paradigm

The avian embryo, as an experimental model, has been of utmost importance for seminal discoveries in developmental biology. Among several approaches, the formation of quail-chicken chimeras and the use of the chorioallantoic membrane (CAM) to sustain the development of ectopic tissues date back to the last century. Nowadays, the combination of these classical techniques with recent in vitro methodologies offers novel prospects to further explore organ formation.
Here we describe a two-step approach to study early- and late-stages of organogenesis. Briefly, the embryonic region containing the presumptive territory of the organ is isolated from quail embryos and grown in vitro in an organotypic system (up to 48 h). Cultured tissues are subsequently grafted onto the CAM of a chicken embryo. After 10 days of in ovo development, fully formed organs are obtained from grafted tissues. This method also allows the modulation of signaling pathways by the regular administration of pharmacological agents and tissue genetic manipulation throughout in vitro and in ovo developmental steps. Additionally, developing tissues can be collected at any time-window to analyze their gene-expression profile (using quantitative PCR (qPCR), microarrays, etc.) and morphology (assessed with conventional histology and immunochemistry).
The described experimental procedure can be used as a tool to follow organ formation outside the avian embryo, from the early stages of organogenesis to fully formed and functional organs.

This article provides an updated approach to the classical quail-chicken chimera system to study organ formation, by combining novel in vitro and in ovo experimental procedures.

The avian embryo, as an experimental model, has been of utmost importance for seminal discoveries in developmental biology. Among several approaches, the ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

Wholemount In Situ Hybridization for Astyanax Embryos

In recent years, a draft genome for the blind Mexican cavefish (Astyanax mexicanus) has been released, revealing the sequence identities for thousands of genes. Prior research into this emerging model system capitalized on comprehensive genome-wide investigations that have identified numerous quantitative trait loci (QTL) associated with various cave-associated phenotypes. However, the ability to connect genes of interest to the heritable basis for phenotypic change remains a significant challenge. One technique that can facilitate deeper understanding of the role of development in troglomorphic evolution is whole-mount in situ hybridization. This technique can be implemented to directly compare gene expression between cave- and surface-dwelling forms, nominate candidate genes underlying established QTL, identify genes of interest from next-generation sequencing studies, or develop other discovery-based approaches. In this report, we present a simple protocol, supported by a flexible checklist, that can be widely adapted for use well beyond the presented study system. It is hoped that this protocol can serve as a broad resource for the Astyanax community and beyond.

Wholemount In Situ Hybridization for Astyanax Embryos

This protocol enables visualization of gene expression in embryonic Astyanax cavefish. This approach has been developed with the goal of maximizing gene expression signal, while minimizing non-specific background staining.

In recent years, a draft genome for the blind Mexican cavefish (Astyanax mexicanus) has been released, revealing the sequence identities for thousands of ...

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in developmental biology for decades. Traditionally, most gene expression studies relied on visual evaluation of the ISH signal, a method that is prone to bias, particularly in cases where sample identities are known a priori. We have previously reported on a method to circumvent this bias and provide a more accurate quantification of ISH signals. Here, we present a simple guide to apply this method to quantify the expression levels of genes of interest in ISH-stained embryos and correlate that with their corresponding genotypes. The method is particularly useful to quantify spatially restricted gene expression signals in samples of mixed genotypes and it provides an unbiased and accurate alternative to the traditional visual scoring methods.

Gene editing technologies have enabled researchers to generate zebrafish mutants to investigate gene function with relative ease. Here, we provide a guide to perform parallel embryo genotyping and quantification of in situ hybridization signals in zebrafish. This unbiased approach provides greater accuracy in phenotypical analyses based on in situ hybridization.

In situ hybridization (ISH) is an important technique that enables researchers to study mRNA distribution in situ and has been a critical technique in ...

Understanding Early Organogenesis Using a Simplified In Situ Hybridization Protocol in Xenopus

Summary

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Chapters in this video

Stem cell-like Xenopus Embryonic Explants to Study Early Neural Developmental Features In Vitro and In Vivo

Functional Cloning Using a Xenopus Oocyte Expression System

Technique to Target Microinjection to the Developing Xenopus Kidney

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

Two-step Approach to Explore Early- and Late-stages of Organ Formation in the Avian Model: The Thymus and Parathyroid Glands Organogenesis Paradigm

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

Wholemount In Situ Hybridization for Astyanax Embryos

Genotyping and Quantification of In Situ Hybridization Staining in Zebrafish