Alkaline Phosphatase in situ assay
Tissue preparation - Frozen sections - AP in situ assay - References
Supplies / Equipment:
- container with liquid nitrogen
- container with dry ice
- long forceps
- freezer (-80°C)
- surgical instruments
- Tissue-Tek Cryomolds
(Disposable Vinyl Specimen Molds (e.g. 25 x 25 x 5 mm), Cat. # 4557
Miles Inc., Diagnostics Division, Elkhardt, IN 46515 USA)
- Tissue Freezing Medium for frozen tissue specimens
(Triangle Biomedical Sciences, Durham, NC, USA
Distributed by Fisher Scientific (Cat. #15-183-13))
- Zip-loc bags
- Aluminum foil
- Fill container with liquid nitrogen (enough to cover all samples).
- Label zip-loc bags and cryomolds.
- Fill cryomold with tissue freezing medium.
- After sacrificing the animal, prepare target tissue as quickly as possible. Place it on the cryomold with interesting side either up or down (sections will be transversal). Make sure sample is covered with tissue freezing medium.
- With the long forceps, place cryomolds onto liquid nitrogen to let freeze from the bottom up while floating. Samples will sink when completely frozen.
- When all the samples are frozen, take them out of the liquid nitrogen, wrap in aluminum foil, put in zip-loc bags, and place in container with dry-ice until finished.
- Freeze all samples at -80°C until making frozen sections.
Supplies / Equipment:
- cryocut (e.g. Cryocut 1800, Reichert-Jung)
- 2 small brushes
- coated slides (e.g. Superfrost Plus, precleaned; will attract tissue sections electrostatically, Fisher 12-550-15)
- Adjust sample to temperature in the cryostat (e.g. -18 to -13°C).
- Cut sections 5 - 8 µm thick. If necessary, flatten with small brush.
- Touch section with slide to pick up tissue section.
Supplies / Equipment:
- three 50 ml Coplin staining jars with glass cover
- wet ice, 1.5 ml Ependorf tubes
- hydrophobic marker (Pap Pen, Kiyota, Japan, from: Ted Pella Inc. #22303)
- xylene-resistant marker or pencil
- humidity chamber
- waterbath at 65°C
- paper towels, gauze sponges (10 x 10 cm)
- coverslips: small (22 x 22 mm) up to large (50 x 24 mm)
- HBHA (Hanks balanced salt solution with 0.5 mg/ml BSA, 0.1% NaN3, 20 mM HEPES [pH 7.0])
- HBS (HEPES balanced saline)
- AP buffer 2 (Alkaline Phospatase buffer: 100 mM Tris-HCl [pH 9.5], 100 mM NaCl, 5 mM MgCl2)
- AP stain (AP buffer containing 0.17 mg/ml BCIP and 0.33 mg/ml NBT)
- Acetone / Formaldehyde fixative (60% acetone, 3% formaldehyde, 20mM HEPES, pH 7.0)
- Turn on 65°C waterbath. For later heat inactivation step, fill Coplin jar with HBS, cover with lid and place in waterbath. Let adjust to 65°C for at least one hour before using.
- Prepare incubation solutions, store on ice. One 50 ml Coplin jar holds 14 slides. Even for small sections, at least 200 µl are needed. To cover all of the slide's surface, 1 ml is usually sufficient. Use two different negative controls: (a) the serum-free cell culture medium (MEM + HEPES for 293 cell c.m.) to check for successful heat inactivation of endogenous AP, and (b) SEAP (secreted alkaline phosphase) containing c.m., to check for binding related to SEAP itself. To calculate the amounts of tagged protein the following estimation could be useful: SEAP: 1000 mU = 1µg, SEAP-PLx: 600 mU ~ 1µg. To achieve good staining, 100 to 500 mU/ml of fusion protein are needed. If you consider, that the label on the c.m. represents mU per 100 µl c.m., the incubation mix should contain at least pure "10 mU" c.m (or 10% "100 mU" c.m.).
- Take frozen sections out of freezer. Let the the white frost dry out (not very time critical, but try to stay under 1 hour)
- Circle tissue sections with hydrophobic marker (helps to save reagents).
- Label slides with xylene-resistant marker or pencil (species, tissue, stage of pregnancy, incubation mix, date).
- Soak frozen sections once (~ 5 min.) in HBHA.
- Drain slides, and wipe dry with gauze the back and the edges of the front of the slides and overlay samples with fusion protein supernatant for 75 min. at room temperature in humidity chamber (moist and level). Do not allow sections to become dry.
- Wash 6 x 5 min. in HBHA + 0.1% Tween 20. First, drain the slide by tapping on paper towels, rinse briefly in first wash and place in second wash. Agitate at the beginning, middle and end of each wash.
- Fix with acetone / formaldehyde fixative for 2 min.
- Wash 3 x 5 min. with HBS (not time critical).
- Heat inactivate endogenous AP for 30 min. at 65°C (transfer slides into Coplin-jar with HBS in 65°C waterbath).
- When time is up, transfer slides to coplin jar with with AP buffer 2 (cool & rinse to pre-adjust pH and temperature).
- Drain slides, and wipe dry with gauze the back and the edges of the front of the slides. Do not allow sections to become dry. Overlay sections with AP stain (NBT + BCIP) for 2 hours (monitor staining on white background). This needed staining time can vary between 10 minutes and 16 hours. When staining is sufficient, drain off stain (toxic !) onto paper towel.
- Rinse 5 x in dH2O.
- Let sections air dry for 30 min. Do not use ethanol / xylene dehydration, since NBT / BCIP will wash out!
- Immediately before coverslipping, rinse slide briefly (< 1 min.) in xylene. Coverslip with Permount. Let slides air dry to non stickyness (usually ~3 days at room temperature).
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