Protocol for Staining Collagen IV

Equipment

Leica CM1950 automatic sectioning cryostat.
Thermo Fisher Scientific round (12 mm diameter) cover glass (thickness No 1.5).
Thermo Fisher Scientific ColorFrostTM Plus slides made of positively charge uncoated glass.

Tissue-tek OCT 4583 compound from VWR.
Tissue-tek cryomolds from VWR.

Urea/Glycine buffer (6M Urea/0.1M Glycine, pH 3.5)

Urea 36g
Glycine 0.75g
Bring to 100 mL with dH₂O, stir to dissolve and adjust pH to 3.5.
Filter sterilize and store @4^oC

NaCl 8 g
KCl 0.2 g
Na₂HPO₄ 1.44 g
KH₂PO₄ 0.24 g
Tween 20 2 ml
Fetal calf serum 50 ml
Bring to 1 L with dH₂O, stir to dissolve and adjust pH to 7.4.
Filter sterilize and store @4^oC

Mowiol 4-88 2.4 g
Glycerol 6 g
dH₂O 6 ml
Stir for 2hrs at room temperature. Add,
0.2 M Tris 12 ml
Heat to 50^oC for 10 min and stir overnight at room temperature.
Centrifuge the solution to remove precipitates at 5,000 x g for 15min; aliquot and freeze the supernatant at -20^oC.

Cutting cryostat sections

1. Wash ColorFrost^TM Plus slides (made of positively charged glass) by dipping in acetone, air dry, and store at room temperature. Store plastic tweezers and brush in the cryostat. Precool cryostat chamber, microtome and a specimen holder (chuck).
   (The use of positively charged glass ensures that sections stick firmly to the slides during the immunolabeling and washing steps.)

2. Rapidly freeze freshly dissected tissues on dry ice. Place Tissue-tek OCT 4583 compound on precooled cryostat chuck and mount the tissue block onto the chuck so that the side of the cut section is at 90^o to the blade. Cut 6-8 μm tissue sections.
3. Place the slide in a precooled slide box and store in a -20^oC freezer. Cut sections are stable for up to 1 month at -20^oC.

1. Take out the slides from the -20^oC freezer, and dip into a jar filled with -20^oC methanol. Fix at -20^oC for 20 min.
2. Let slides air-dry for 20min. Dip slides into a jar filled with Urea/Glycine buffer. Fix at room temperature for 30min.
3. Wash slide 3 times with 1x PBS buffer.
4. Make a wet chamber by lining a 24 x 24 cm petri dish with two or three sheets of filter paper and add water to moisten the filter paper.
5. Wash slides once with BLOCK solution. Drain BLOCK solution off against a towel and put slides into the wet chamber.
6. Dilute primary antibody (anti-collagen) at 1:100 in BLOCK solution. Add the primary antibody solution to each slide (300-500 μl is sufficient to cover the section on each slide).
7. Incubate the wet chamber at 4^oC for overnight.
8. Take slides out of the wet chamber, drain off antibody, and wash three times with the BLOCK solution.
9. Dilute secondary antibodies (anti-rabbit conjugated to FITC, TRITC or other fluorophores) at 1: 200 in BLOCK solution. Add the secondary antibody solution to each slide and put slides back into the wet chamber.
10. Incubate the wet chamber at 4^oC for overnight.
11. Take slides out of the wet chamber, drain off antibody, and wash three times with the BLOCK solution.
12. Drain BLOCK solution off against a towel. Add 20 μl of Mowiol solution to the section on each slide. Place a cover glass onto the section and apply a small pressure to remove any trapped air bubble.
13. Let Mowiol solution dry at room temperature for 2 hours. Store glass slides at 4^oC for no longer than 1 month.