Studying Wnt Signaling During Patterning of Conducting Airways

Animals were housed in pathogen-free conditions. Mice were handled according to protocols approved by CCHMC Institutional Animal Care and Use Committee (Cincinnati, OH USA). Mice utilized throughout these studies were maintained in a mixed background.

1. Whole Mount X-galactosidase Staining

1. Euthanize pregnant female at E11.5 to E14.5, by CO₂ inhalation. Place animals in CO₂ chamber, charge the chamber with CO₂. Maintain animals in chamber for minimum of 5 min. Perform secondary method of euthanasia by cervical dislocation.
2. Clean abdominal area with ethanol (EtOH) 70% and perform laparotomy with a single abdominal midline incision. Utilize surgical scissors and standard pattern (straight serrate) forceps.
3. Isolate uterine horns using forceps and fine scissors and immediately place tissue in petri dish containing chilled PBS solution. Maintain tissue on ice until proceeding to isolation of embryos.
4. Isolate embryos from uterine horns. Dissect embryonic tracheal lung tissue using dissecting microscope. Utilize fine tip forceps and scissors to hold and puncture the uterine tissue and concepti releasing the embryos.
   1. Remove remaining embryonic membranes. With assistance of needle blades cut head of embryos and lower body part below the diaphragm. Locate liver as a landmark.
   2. After isolation of the thoracic region of the embryo, proceed to remove lateral sides of the body wall, spine and heart using needle blades. Take care while removing the heart to avoid injuring tracheal tissue. Finally, carefully, separate the esophagus from the trachea by pulling the esophagus with assistance of needle blades. Maintain tissue in PBS solution on ice.
5. Place tissue in 4 ml screw cap vials using glass or transfer pipettes. Fix tracheal lung tissue in 4% paraformaldehyde (PFA) in PBS for 30 min. After fixation, rinse tissue with PBS. While rinsing, prepare X-gal staining solution (add 5 mM K₃Fe(CN)_6, (100 μl 0.5 M stock) 5 mM K₄Fe(CN)₆ (100 μl 0.5 M stock) 2 mM MgCl_2, (20 μl 1 M stock) 0.01% NaDOC (50 μl 2% stock), 0.02% NP₄O (100 μl 2% stock), 1 mg/ml X-gal (500 μl of 20 mg/ml stock) and 9.13 ml of distilled water to bring to a final volume of 10 ml).
6. Remove any remaining PBS and add 2 ml of X-gal staining solution to the glass vial. Place vials in tray on shaker. Stain tissues 1 to 2 hr in X-gal staining solution. Incubate the tissue in X-gal solution for 4 hr for further processing and embedding.
7. Stop the reaction by washing tissues in 3% dimethyl sulfoxide-PBS. Rinse in PBS, 3 times for 5 min each wash and store in 70% ethanol.
8. Fill Petri dishes with a 1% agarose/PBS solution. Allow agarose to solidify. Using glass transfer pipettes, place the tissue on agarose coated plates filled with PBS solution. Photograph whole mounts using a dissecting microscope. Select appropriate filter for brightfield.
9. To better distinguish sites of X-gal staining process samples for paraffin embedding and sectioning. Place samples in fine screen cassettes.
   1. To process samples for paraffin embedding use an automated processor and set up a short cycle as follows: six changes of alcohol: 5 min first change and 3 min per each remaining change. Three changes of Xylene: 5 min first change and 3 min next two changes. The previous steps are performed at 30 °C. Finally, perform three changes of paraffin at 62 °C: 25 min, 8 min and 5 min.
10. Orient samples for embedding with tissue lying flat on the bottom of embedding boat. Carefully add paraffin to fill the embedding boat. Cool immediately on cold plate of embedding station until paraffin solidifies. Remove block from embedding boat. Using a microtome, generate 6 μm sections. Mount sections on pretreated-slides and dry slides on slide warmer set at 42 °C for 1 hr.
11. Bake slides overnight in oven at 56 ºC. Place slides in racks. Deparaffinize slides with three changes of 100% Xylene, 10 min each. Use staining dishes filled with 200 ml of Xylene.
12. Rehydrate slides using graded EtOH series (100%, 70%, 50% and 30%, 1 min per step) to PBS before counterstaining with Nuclear Fast Red for 10 to 30 sec. Rinse slides with tap water three times for 5 min each time to remove excess of Nuclear Fast Red.
13. Dehydrate slides in a graded EtOH series (50%, 70% and 100% EtOH) before washing in three changes of Xylene (20 dips each change) and cover slipping with Xylene based mounting media.

2. Lectin Staining

1. Dissect E13.5 and E14.5 tracheal lung tissue as described in steps 1.1 to 1.4. Using glass transfer pipettes place tissue in screw cap vials. Fix tracheal lung tissue with 2 ml of 2% PFA solution overnight at 4 °C. After fixation, rinse samples in PBS three times for 10 min each wash.
2. Prepare blocking buffer, usually a final volume of 10 ml of solution. Add 0.1 g of BSA (1%) to 8 ml of PBS. Allow BSA to dissolve. Add 0.2 ml of goat serum (2%) 0.03ml of Triton X-100 (0.3%). Bring final volume to 10 ml with PBS. Block samples for 1 hr in 0.5 ml of blocking buffer at room temperature.
3. Remove blocking buffer using a transfer pipette and replace with lectin solution comprised of 50 μg/μl of lectin, 10% goat serum and PBS. Use 250 μl of lectin solution per sample. Incubate overnight at 4 °C. Make sure to cover the glass vial containing tissue with aluminum foil. After incubation with lectin solution, rinse samples in PBS three times for 10 min.
4. Place explants in agarose coated petri dishes containing enough PBS to cover samples. Photograph whole mounts using a fluorescence-dissecting microscope. Select the appropriate filter to detect fluorescence. Lectin PNA is usually coupled to GFP.

3. Whole Mount Immunofluorescence Staining and Confocal Microscopy

1. Isolate tracheal lung tissue, transfer to 4 ml glass screw cap vials and fix overnight in 4% PFA. E11.5 tissue can be fixed in 2% PFA. Store in methanol 100%. Samples can be stored up to several months at -20 °C.
2. Remove methanol using transfer pipette and permeabilize samples using Dent's Bleach (4 parts Methanol, 1 part DMSO and 1 part 30% H₂O₂) for 2 hr. Hydrate tissue in a graded series of methanol diluted in PBS as follows: 100% methanol, 75% methanol, 50% methanol, 25 % methanol, 100% PBS. Incubate 10 min at room temperature during each step of hydration.
3. Block tissue in 0.5% blocking reagent diluted in PBS (see Materials for details) for two hours at room temperature with agitation.
4. Dilute primary antibody in blocking solution (Sox9 1:100, Nkx2.1 1:200, αSMA 1:250) and incubate samples overnight at 4 °C. Wash samples with PBS five times at room temperature. Perform each wash for 1 hr.
5. Apply secondary antibody at a 1:500 dilution in 0.5% blocking solution. Carefully select secondary antibodies to prevent binding among secondary antibodies. Incubate overnight at 4 °C in dark room. Wash samples three times for 20 min at room temperature. Dehydrate samples in a graded series of methanol diluted in PBS as follows: 25% methanol, 50% methanol 75% methanol, 100% Methanol, 10 min each step.
   NOTE: Ensure that samples remain covered with aluminum foil and minimize exposure to light. Tissue can now be stored at 4 °C for several weeks until photographed.
6. Transfer samples to custom made stage plates, remove methanol and clear with approximately 200 μl of Murray's clear solution^19,20 (2 parts Benzyl benzoate, 1 part Benzyl alcohol) immediately before imaging. Photograph using a confocal microscope. Process and analyze image using imaging software.

4. Cell Proliferation

1. Inject E11.5 pregnant mice intra-peritoneally with a solution of BrdU at a concentration of 100 μg BrdU/g of body weight. Sacrifice female as described in section 1 of the protocol and isolate embryos at E12.5 or E13.5. Using knife blades, excise heads and lower part of body below thoracic cavity.
2. Place thoracic tissue in 4 ml screw cap vials and fix in 1 ml of 4% PFA overnight. Wash samples with PBS twice for 5 min and dehydrate samples through an Ethanol series diluted in distilled water up to 70% ethanol as follows: 30%, 50% and 70% ethanol, for 5 min each step.
   1. Place samples in medium size screen cassettes. Process tissue for paraffin embedding using and automated processor. Set up a cycle as follows: six changes of alcohol for 8 min each, three changes of Xylene for 6 min each, three changes of paraffin, for 25 min, 9 min and 8 min.
3. Embed in paraffin orienting the tissue in desired position to generate 6 μm transverse or longitudinal sections as described in section 1. Place sections on slides and allow tissue to adhere to slide on slide warmer set at 42 °C for 1 hr.
4. Bake slides, on their sides, overnight in oven at 56 °C. Place slides in plastic racks. De-paraffinize slides by immersion in three changes of 100 % Xylene, 10 min per change. Use 200 ml of Xylene to completely submerge slides. Rehydrate slides using graded EtOH series (100%, 70%, 50% and 30%, 5 min per step with agitation) to PBS (5 min).
5. Perform antigen retrieval using 10 mM citrate buffer, pH 6. To prepare 100 ml of citrate buffer mix 18 ml of 0.1 M Citric acid monohydrated and 82 ml of 0.1 M Sodium citrate.
   1. Place samples in plastic coplin jars filled with antigen retrieval solution and microwave for 6.5 min at high power. Add distilled water to coplin jar to compensate for evaporated-citrate buffer and reheat at 40% power for 6 min.
   2. Fill coplin jar to top with water, and reheat again at 40% power for 6 min. Microwave times may vary based on microwave used.
   3. Allow slides to cool for 20 min at room temperature before washing slides in distilled water for 1 min, followed by PBS for 5 min. After the PBS wash, block slides for 2 hr in blocking solution containing TBS (2.425 g of Tris base and 8.765 g of NaCl diluted in 1 L of distilled water) 10% Donkey serum and 1% BSA.
6. Add primary antibodies diluted in blocking solution (TBS containing 10% Donkey serum and 1% BSA). BrdU (mitosis marker), Nkx2.1 (respiratory tract epithelium), Sox9 (cells that will give rise to cartilage) and αSMA (smooth muscle cells). Incubate slides overnight, at 4 °C. Use overlay method to conserve antibodies. Apply 180 μl of antibody to a gasket and then attach slide as if cover slipping.
7. Detach gasket by dipping slides into distilled water. After removal of the gasket, wash unbound primary antibody by performing six 5 min washes with TBS containing 0.1% Tween-20.
   1. Incubate slides with secondary antibody diluted in blocking solution at a dilution of 1:200, for 1 hr at room temperature. Select secondary antibodies carefully to prevent undesired binding among them. Remove unbound secondary antibody by performing three five minute washes with TBS containing 0.1% Tween.
   2. Wash slides in 0.1 M Tris base twice for 5 min and then 0.05 M Tris base twice for 5 min. Slides may be left in 0.05 M Tris base while coversliping using mounting media with or without DAPI (1.5 μg/ml). Store slides in slide folder at 4 °C and protect from light by covering folder with aluminum foil.
8. Visualize staining and photograph using an automated fluorescence microscope. Count labeled-cells and total cells per field photographed at 20X and 40X to determine ratios of proliferating cells to total cells.