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

1. Embryo Preparation

1. 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.
   1. 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.
2. 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.
   1. 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 °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.
   2. 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 °C for 1-2 weeks.
   3. 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 °C as stocks.
   4. 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 °C.
   5. 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.
   6. Store a solution of 100% methanol at -20 °C. After the paraformaldehyde fixation, replace the Mempfa solution with approximately 4 ml of the -20 °C, 100% methanol for storage of the embryos.
   7. 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.

2. Probe Preparation

1. Use 1-2 µg of template DNA to make the digoxigenin-labelled probes. Cut plasmid containing the appropriate DNA sequence at the 5’ 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).
2. 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 µl. Assemble the reaction at room temperature because components of the polymerase buffer can precipitate the template DNA when in high concentrations and cold.
3. Incubate the transcription reaction for 2 hr at 37 °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.
   1. 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 µg 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.
   2. Add 2 µl ethidium bromide stock solution (10 mg/ml) to about 100 ml of agarose gel when the agarose has cooled to about 60 °C to allow visualization of RNA under ultraviolet (UV) light.
      NOTE: This agarose gel solution can be kept in a 60 °C incubator so that future gels can be poured without repeated melting. Take care in handling ethidium bromide due to potential toxicity.
4. Add 1 µl DNAseI (RNAse free grade) to the transcription reaction after the 2 hr incubation and incubate for a further 10 min at 37 °C to eliminate template DNA.
5. Remove 1 µl of the reaction mix to check on the 1% agarose-TAE gel and to the rest (20 µl) add 80 µl of 1% SDS in TE buffer (10 mM Tris pH 8.0, 1 mM EDTA), 10 µl of 5M NH₄ Acetate, and 220 µl of cold ethanol. Vortex the mix vigorously and set aside on ice until the results of the RNA quality check are known.
   1. Run the 1 µl of RNA removed before precipitation on the 1% agarose-TAE gel. To ease loading on the gel, add 4-5 µl of water and 1 µl 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).
6. 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.
   1. Resuspend the probe with 1 ml of RNA hybridization buffer (Table 1) in the eppendorf tube.  Vortex and briefly heat the tube to 37 °C and vortex again.  Transfer the probe solution to a 15 ml screw cap polystyrene tube and fill to 7-10 ml with RNA hybridization buffer.  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.

3. In situ Hybridization

1. Take the embryos that were stored in -20 °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.
   1. 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.
   2. Once in the probe solution, hybridize the embryos overnight as described in Table 3.
2. 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 °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.
   1. 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.
   2. 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.
3. 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.
   1. Replace the last wash solution with the BM Purple alkaline phosphatase substrate. Perform the staining reaction at either room temperature or 37 °C.
      NOTE: Staining is more rapid at 37 °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.
   2. 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 °C. If the target mRNA is very abundant, put the embryos in the staining solution at 4 °C overnight and then move the embryos to room temperature or 37 °C to allow better monitoring of the staining reaction.
   3. 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.
4. 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 °C freezer.
   1. Rehydrate the embryos and fix the stain with Mempfa (Table 6). Once fixed, remove the Mempfa and wash the embryos with 25% methanol.
   2. 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).
   3. 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). 

4. Imaging Embryos

1. 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.
   1. 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 °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.
   2. 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.
   3. 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.
2. 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.
3. 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.
   1. 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.
   2. 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.
   3. 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.

5. Double In situ Hybridization

1. 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.
   1. 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.
   2. 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.
2. 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.
   1. 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.
3. 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 °C overnight.
   1. The following day, wash the embryos thoroughly in MAB (12 washes of 30 min) to remove the excess antibody.
   2. 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).
   3. 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.
   4. 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.