Materials 21 Tissue and Slide Preparation

1. Cryostat (a rotary microtome set in a freezer).

2 Isopentane: Place in a stainless-steel beaker set in an ice bucket filled with a dry ice/acetone mixture

3. Black boxes:

a. Plain plastic boxes for 25 slides: These protect against condensed moisture for freezer storage of slides.

b. Black boxes with 25-slide polyacetyl rack and dessicant compartment: These are used for exposure of slides to the emulsion.

4. Tissue culture standard "Milli Q" purified water (i e., filtered through activated charcoal, deionizing resins, and an ultrafilter).

5. All solutions used on the tissue prior to hybridization should be made with diethyl pyrocarbonate (DEPC; Sigma, St Louis, MO) treated water. Add 1 mL of DEPC to 1 L of "Milli Q" H20, and shake vigorously Allow to stand several hours or overnight, and then autoclave

6. Fixatives:

a. 4% (w/v) Paraformaldehyde in 0.1Mphosphate buffer pH 7.4: Heat 4 g paraformaldehyde in 50 mL deionized water to 60-70°C and titrate until clear with 10N NaOH (a few drops). Cool to 40°C, and then mix with 50 mL of 0.2M phosphate buffer.

b. Picric acid-paraformaldehyde (PAF) in 0.1M phosphate buffer pH 7.4: Heat 4 g paraformaldehyde in 30 mL of saturated picric acid to 60-70°C, and titrate until clear with 107V NaOH. Cool to 40°C. Mix with H20 to a volume of 100 mL, and then add an equal volume of 0.2M phosphate buffer.

Na2HP04 (28.2 g/1) NaH2P04 (6.9 g/250 mL) Titrate the above two solutions to pH 7.3. Treat with DEPC (0.1% [v/v]), and then autoclave.

9. PBS containing heparin is the initial wash for the perfusion start. To 100 mL of 20 mM phosphate buffer diluted from the above stock with DEPC-treated water, add 8 g sodium chloride and 100 U of heparin

10. 30% (w/v) sucrose in QAM phosphate buffer.

11. Perfusion fixation setup (optional; see Note 2):

a. Two 1-L bottles with hose connections at the bottom.

b. Silicone tubing (3/g in.) to fit the bottle hose connections.

c. Three-way stopcock, which fits the above tubing.

d. "Venoset" from a medical supply distributor provides a bubble trap, tube, and tubing clamp, and a louer slip end, which fits any syringe needle The top will fit the 3/8-in. tubing connected to the stopcock.

f. "Intramedic" polyethelene tubing (the largest size that will fit into the aortic artery of the animal—for rats 7430, for mice 7405). Take a piece about 1.5-in. long, hold it up to a match flame, and flare one end. Fit the other end onto a tight-fitting needle.

Place the bottles 75 cm above the animal. This provides adequate pressure to perfuse rats and mice. Fill one bottle with fixative, and let it flow through to the stopcock. Fill the other bottle with an isotonic buffered saline (e.g., PBS), and fill the tubing with the bubble trap upside down until the trap is half full. Turn the trap leaving it half full, fill the remainder of the tubing, and close the clamp The catheter is inserted according to the directions given below with the clamp slightly open allowing a slow flow. There must be a positive pressure on the catheter tip as it is inserted to avoid any air bubbles entering the blood vessels. Air will block the flow. Once the tip is in place increase the flow.

12. Microslides, precleaned Superfrost (Dynalab, Rochester, NY or Fisher Scientific, Pittsburgh, PA). These do not have to be washed prior to use. Many other brands first have to be washed with detergent.

13. Acid cleaning solution (for cleaning slides and glassware):

Potassium dichromate, 20 g.

Water, 200 mL.

Sulfuric acid, 20 mL

14. 0.25M Ammonium acetate.

15. 50 |ig/mL Poly-l-lysine (Sigma #P1274) in 10 mM Tris-HCl buffer (pH 8.0).

Stock solutions:

10 mM Tris-HCl buffer pH 8.0.

Poly-l-lysine 10 mg/mL in 10 mM Tris-buffer. pH 8.0. Store at -20°C in 1.5-mL aliquots.

Dilute 200X to make the working solution (1.5 mL in 300 mL 10 mM Tris-HCl).

16. Glass staining jars, acid cleaned.

17. Slide racks to fit staining jars (may be glass or metal).

18. OCT (a "Tissue Tek" product, Miles Inc., Elkhart, IN) for mounting small frozen tissue pieces to microtome chuck. Sterile deionized water can also be used and is preferable with fresh frozen tissue, since the OCT does not wash away after fixation in paraformaldehyde and the probes have a tendency to stick to it, thereby increasing the background.

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