The overall goal of the staining procedures is to demonstrate the role of the Wnt signaling pathway in the early stages of development of the upper airways using embryonic tissue from genetically engineered mouse models. This methods can help answer key questions in the field of pulmonary biology. Such as, for example, how the supporting structures of the conducting airways are patterned.
The techniques that we're going to presenting here will allow us to study cell proliferation or the process of mesenchymal condensations, which are key events in the formation of the tracheal cartilage. The main advantage of this technique is it can be used on small amounts of tissue. This includes lung explants, and tracheas isolated from E11 embryos.
Generally speaking, individuals new to this procedure will struggle do to the size of the sample, as well as a limited amount of tissue available. One must feel comfortable when working with embryonic tissues, and will do so by paying attention throughout the steps and making sure that the same amount of tissue is available from step to step. After euthanizing a pregnant female mouse at E11.5 to E14.5, and performing a laparotomy according to the text protocol, use forceps and fine scissors to isolate the uterine horns, and immediately place the tissue into a Petri dish containing chilled PBS solution.
Then place the dish on ice. To isolate the embryos from the uterine horns, use fine forceps to hold the uterine tissue. And with scissors, puncture the uterine tissue and conceptus, releasing the embryos.
Next, remove any remaining embryonic membranes. Then, under a dissecting microscope using needle blades, cut off the head and lower body, below the diaphragm, and locate the liver as a landmark. Use needle blades to remove the lateral sides of the body wall, spine, and heart.
Taking care not to injure the tracheal tissue. To dissect the embryonic tracheal lung tissue, carefully separate the esophageus from the trachea by using needle blades to pull the esophageus. Place the tissue in PBS on ice before using glass or transfer pipettes to transfer the tissue into four millimeter screw cap glass vials.
Add 4%paraformaldehyde, or PFA, in PBS and incubate for 30 minutes. Then use PBS to rinse the tissue. After removing the PBS from the samples, add two millimeters of X-gal staining solution and incubate for one to two hours, or four hours for further processing and embedding.
Stop the reaction by using 3%DMSO in PBS to wash the tissue three times for five minutes each. Then store the tissue in 70%ethanol. After using an automated processor to process the samples for paraffin embedding according to the text protocol, orient the samples with the tissue flat on the bottom of an embedding boat.
Carefully add paraffin to fill the embedding boat, and immediately place on the cold plate of the embedding station until the paraffin solidifies. Next, remove the block from the embedding boat, and with a microtome, cut six micrometer sections. Then mount the sections on slides.
And place the sections on a slide warmer at 42 degrees Celsius for one hour. Bake the slides in a 56 degree Celsius oven overnight. Then, to deparrafinize the samples, place the baked slides in racks and transfer into staining dishes filled with with 200 milliliters of 100%xylene, three times for 10 minutes each.
Use a graded ethanol series for one minute each to rehydrate the slides. Then use Nuclear Fast Red to counterstain the samples for 10 to 30 seconds. With tap water, rinse the slides three times for five minutes each.
To dehydrate the slides, use a graded ethanol series before washing the samples in three changes of xylene. Then use a xylene-based mounting medium and cover slips to mount the samples. To carry out lectin staining, after fixing tracheal lung tissue and preparing blocking buffer according to the text protocol, transfer samples to 0.5 milliliters of blocking buffer and incubate at room temperature for one hour.
Use a transfer pipet to remove the blocking buffer and replace with 250 microliters of lectin solution per sample. Then using aluminum foil, cover the glass vials, and incubate at four degrees Celsius overnight. The following day, after using PBS to rinse the samples, place the explants in agarose-coated Petri dishes containing enough PBS to cover the samples.
Then use a florescence dissecting microscope to photograph the whole mounts. To perform immunofluorescence staining on whole mounts, after fixing the samples according to the text protocol, use a transfer pipette to remove the methanol, then add dense bleach and incubate the tissue for two hours to permeabilize the samples. Using a graded series of methanol, diluted in PBS, hydrate the tissue by incubating it at room temperature for 10 minutes at each step.
After blocking the tissue according to the text protocol, add primary antibodies and blocking solution, and incubate the samples at four degrees Celsius overnight. Once the samples have been washed in PBS, add a one to 500 dilution of secondary antibodies in PBS, and incubate in the dark at four degrees Celsius overnight. The following day, after washing the samples three times for 20 minutes each in PBS, use a graded methanol series in PBS to dehydrate the samples for 10 minutes at each step.
Transfer the samples to custom-made stage plates. Remove the methanol. Then use 200 microliters of Murray's clear solution to clear the samples.
Image immediately using a confocal microscope. After injecting E11.5 pregnant mice with BrdU, and isolating the embryos at E12.5 or E13.5 according to text protocol, use knife blades to excise the embryonic heads and lower parts of the bodies, below the thoracic cavities. Place the thoracic tissue in four milliliter screw cap vials, and add one milliliter of 4%PFA.
After embedding, sectioning, and rehydrating samples according to text protocol, place the slides in Coplin jars and fill with 10 millimolar citrate buffer, pH six. Microwave the samples at high power for six and a half minutes before using distilled water to replace the evaporated citrate buffer and heating again. After cooling, washing, and blocking the slides according to the text protocol, apply 180 microliters of primary antibodies, diluted in blocking solution, to a gasket and attach the slide as if applying a cover slip.
The following day, dip the slides in distilled water to detach the gasket. Then use TBS, with 0.1%Tween 20, to wash the slides six times for five minutes each. After incubating the slides with secondary antibodies, use 0.1 molar Tris Base to wash the slides twice for five minutes each, before washing twice with 05 molar Tris Base.
Keep slides in 05 molar Tris Base while adding mounting medium, with or without DAPI, and add a cover slip. Finally, use an automated florescence microscope to image the samples, and analyze according to the text protocol. In this figure of whole-mount tracheal lung tissue, isolated from Reporter and two lacZ mice, lacZ staining was detected in the mesenchyme of the trachea, and the peripheral regions of developing lungs, indicating Wnt beta catenin activity.
In this experiment, E12.5 to E14.5 tracheal lung tissues were stained with PNA lectin. In control animals, florescence was detected as periodic bands in mesenchyme where cartilage forms. However, the lack of staining in Wntless sonic hedgehog Cree embryos, indicates that Wnt signaling is necessary for mesenchymal condensations.
In whole mount tracheal tissue of E11.5 embryos, SOX9 expression is limited to mesenchyme as a continues stripe, while in the developing lung, SOX9 is observed at the periphery of the respiratory epithelium. As shown here, NKX2.1 is restricted to the epithelium of the trachea. Prevention of secretion of Wnt ligands from epithelium diminishes expression of SOX9 in mesenchyme, but does not effect SOX9 expression in peripheral lung epithelium.
Alpha smooth muscle actin staining is detected at low levels in tracheal tissue, but it is clearly observed in larynx and stomach. Finally, as indicated in this figure, in vivo labeling of tracheal tissue with BrdU, SOX9, and alpha smooth muscle actin, demonstrates that SOX9 stained cells undergo proliferation at a higher level than alpha smooth muscle actin stained cells. The proliferation pattern is reverted in Wntless sonic hedgehog Cree tracheal tissue.
Once mastered, most of these techniques can be done in two days if performed correctly. While attempting this procedure, it is important to remember the source of the primary antibody when choosing the secondary antibody. This is important for all immunostainings, but it is especially important when you have more than one primary antibody.
Following the X-galactosidase staining, additional techniques such as in situ hybridization can be performed to acquire additional information, as well as answer additional questions, such as whether or not a particular gene of interest will be expressed during Wnt signaling. The techniques presented here would be useful for researchers that are interest in a silent patterning mechanisms mediated by Wnt signaling, or other key signaling powers to development. After watching this video, you should have a good understanding of how to study differentiation process during the development of the respiratory tract.
And also have a better understanding of the roll of Wnt signaling in the patterning of the conducting airways. Don't forget when working with paraformaldehyde and benzyl benzoate to take the proper precautions, like earing gloves, preparing solutions under the hood, and disposing of waste solutions in accordance to your local laws and regulations.
