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

The overall goal of the following experiment is to show where a particular gene is expressed within an intact embryo. This is achieved by fixing the RNA to be localized in place within the embryo as a second step. An antisense RNA probe specific to the gene of interest is hybridized to the target RNA within the embryo.

This probe contains an epitope, usually digo genin, which can then be detected. Next, an antibody coupled to alkaline phosphatase is hybridized to the epitope on the RNA probe, followed by use of a colormetric stain. In order to visualize where the RNA is localized within the embryo results are obtained that show precisely where the gene of interest is expressed based on the color reaction and knowledge of the embryo anatomy.

The main advantage of our of our institute protocol over existing protocols is that it is faster and utilizes fewer reagents with no apparent loss of quality. This method can answer key questions in embryology, such as how tissues are specified and patterned. Though this method can provide insight into the development of opus, it can also be used for many other small embryos, such as zebrafish and early mouse embryos.

We first started developing these methods because the original methods occupied a great deal of time, and we started experimenting with leaving Ed steps to increase the efficiency of the procedure without compromising quality. Embryo fixation is performed in stages. First, prepare the glass vials that will be used for all steps, including the final storage of embryos to allow for monitoring the embryos during all steps of the process.

Use clear vials with a good Teflon seal in the lid using a permanent marker label the vials with the appropriate experimental information. Cover the label with clear tape to prevent the loss of the labeling over the course of the procedure. Due to the use of alcohols and other solvents, prepare the MFA fixation solution as described in the protocol text.

Fill each labeled glass vial with three to four milliliters of the MFA solution. Store a solution of 100%methanol at minus 20 degrees Celsius. Make a cut glass pipette and use it to transfer embryos to the glass vials containing Memphis solution for fixation.

Add the embryos with a minimum of liquid transfer from the embryo medium. Fix no more than 20 to 30 embryos per vile. Leave the embryos in Memphis solution for two hours at room temperature on a mutator.

At the completion of the paraform aldehyde fixation, replace the Memphis solution with approximately four milliliters of the minus 20 degrees Celsius. 100%methanol for storage of the embryos prior to assembling the probe synthesis reaction, allow the DNA water NTP mix and polymerase buffer to warm to room temperature. One of the most critical steps for the success of this procedure is to assemble the probe reaction correctly.

Assemble the probe synthesis reaction at room temperature. Add the following components to a 1.5 milliliter micro centrifuge tube in this order. Template DNA water adjusted to a volume that brings the total volume of the reaction to 20 microliters NTP mix, RNA inhibitor, RNA polymerase buffer and RNA polymerase incubate the transcription reaction for two hours at 37 degrees Celsius.

Incubations slightly longer than two hours, have no adverse effects, but also show little increased yield if incubated for one hour, the yield will be reduced, although still sufficient to make a good probe. When the transcription reaction is complete, remove one microliter of the reaction mix for checking on a 1%aros. TAE gel to the remaining reaction mix.

Add 80 microliters of 1%SDS in TE buffer, 10 microliters of five molar ammonium acetate and 220 microliters of cold ethanol. Vortex the mix vigorously and set it aside on ice until the results of the RNA quality check are known. After adding water and standard loading dye to the one microliter of RNA removed before precipitation, run the RNA on a 1%aros, TAE gel containing AUM bromide.

View the atherium bromide labeled RNA on the gel using a UV light transluminator in order to check the quality of the probe. A single band without smearing either above or below it should be visualized. Once the quality of the RNA has been verified, precipitate the remaining RNA that was set aside on ice by spinning in a micro centrifuge at full speed for 10 to 15 minutes.

Remove the supernatant with a drawn out glass pipette and allow the RNA to dry briefly. Subsequently, the probe is resuspended in RNA hybridization buffer and stored in a 15 milliliter screw cap polystyrene tube at minus 20 degrees Celsius. To begin this procedure, retrieve the embryos that were stored in minus 20 degrees Celsius methanol and allow them to warm to room temperature.

The entire procedure is performed in the glass vials. If different embryos from one group are going to be examined 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 in preparation for the addition of the probe, rehydrate the embryos through a methanol series as outlined in this table, use glass pipettes to remove and add each solution carefully as the embryos can be easily damaged during this step.

When transferring the liquids, check the inside of the caps to ensure that embryos have not been trapped there. Gently swirl the embryos after each change to ensure that they are not sticking to the sides or each other. And rock the embryos by mounting the vials on a mutator prior to use the RNA hybridization buffer and the probe must be pre-warned to 65 degrees Celsius.

Hybridize the embryos in the probe solution overnight with rocking at 65 degrees Celsius on the following day. Remove the probe solution and save it in a 15 milliliter screw cap polystyrene tube marked with the date and number of times the probe has been used. Store the probe solution at minus 20 degrees Celsius.

Once the probe is removed, prepare the embryos for antibody staining against the probe through a series of washes as outlined here, change the temperature as required by moving the mutator with vials of embryos attached directly into hybridization ovens that are set to the appropriate temperature. During the time when the embryos are being incubated in the blocking solution, make up the appropriate volume of the blocking solution plus anti-D dig antibody so that the antibody is blocked prior to its addition to the embryos. Incubate the embryos in the antibody solution overnight with rocking at four degrees Celsius on the following day.

Remove the antibody solution and wash with mab, a solution that contains maleic acid and sodium chloride. Perform at least 1230 minute washes at room temperature with rocking. Replace the last wash solution with the alkaline phosphatase substrate.

Perform the staining reaction at room temperature with rocking overnight on the following day. Stop the staining reaction and prepare the embryos for storage and imaging by changing the liquids as outlined in this table. After rehydrating the embryos, fix the stain with MFA for 30 minutes.

Once fixed, remove the MFA and wash the embryos with 25%methanol. If removal of endogenous pigment is required, remove the 25%methanol and add the bleaching solution. Observe the bleaching closely as it happens relatively quickly, and the degree of bleaching can be varied for different effects.

For long-term storage, dehydrate embryos through a methanol series to 100%methanol following bleaching for short-term storage and subsequent imaging, transfer the embryos to PBS Uncleared embryos are best viewed and imaged on a 1%aros background. To prepare the aros solution, add aros to water and bring to a boil until the aros is in solution. After the aros has cooled to 50 degrees Celsius, pour it into a Petri dish to a depth of about two millimeters.

When the aros dish is ready, place the uncleared embryos in the dish and proceed with imaging. The aros gives a blue gray 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 to image the embryo from alternate, such as from the ventral side.

Use fine forceps to cuts in the aros to fit the embryo. Place the embryos in those channels for orientation. Take care in manipulating the embryos as they are easily damaged.

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. Cleared embryos are used to image staining that is deep within the embryo, such as in the no cord, lung, or regions of the brain.

In this protocol, different levels of bleaching are used to visualize staining in the embryo. If the stain is strong, a lighter bleaching that leaves some pigmentation enables better staging and orientation of the embryo. However, if the stain is in an area with high levels of pigmentation such as the kidney, greater bleaching gives a better result.

Manipulation of the ARO base can help in embryo orientation. For example, by cutting a fine channel in the aros, an embryo at the tad pole stage that is lying on its side can be viewed from the ventral side with enough stability to capture a good image. This embryo was placed in a small hole that stabilized its position with the ventral side up, allowing full viewing of the expression pattern of the gene of interest.

Internal organs can be viewed in both uncleared and cleared embryos in an uncleared opaque embryo stained for PAX two. At stage 34, the optic stalk as well as the neural tube can be visualized. However, details are not sharp By clearing the embryo, the boundaries of expression sites, including the optic stalk are sharper.

The extent of clearing is shown by the ability to see both eyes in this embryo. However, a common problem with cleared embryos is that some staining accumulates in the internal cavities. After watching this video, you should have a good idea of how to perform a whole mount in situ quickly with fewer reagents and capture high quality images.