It has been demonstrated that frozen tissue sections are particularly suitable for RNA in situ hybridization, immunocytochemistry and enzyme histochemistry. The major advantages of cryosectioning over traditional paraffin wax sectioning are that tissue embedding and sectioning are carried out under frozen conditions, and that activities of DNase, RNase and protease are greatly inhibited. Therefore, potential degradation of DNA, RNA or proteins is minimized, which is essential to in situ hybridization and enzyme histochemistry. For this reason, the present section describes in detail the protocols for tissue freezing, specimen preparation, sectioning and fixation.











FIGURE 12.1 Diagram of section ribbons with a trapezoidal shape and mounting on a coated glass slide.

FIGURE 12.1 Diagram of section ribbons with a trapezoidal shape and mounting on a coated glass slide.

Preparation of Frozen Specimens

1. Chill all tools employed for cryosectioning in the cryostat chamber for 15 min, including razor blades, brushes and Pasteur pipettes. However, glass slides should not be prechilled.

2. Prechill a 50-ml Pyrex beaker or its equivalent in a container filled with liquid N2 for 1 min and add a sufficient volume of CryoKwik to the beaker.

3. Label one end of a filter paper strip (3MM Whatman) with a pencil according to the specific sample and carefully place a specimen on the other end of the strip with forceps or a Pasteur pipette.

4. Carefully immerse the specimen in cold CryoKwik for 1 min and then transfer the specimen strip into a cryostat chamber. Alternatively, the specimen at this stage can be stored at -80°C until use.

5. Place a thin layer of OCT compound on the cutting chucks and place the specimen end of the strip onto the layer of OCT compound.

6. Cool the specimen/chuck mount within the cryostat with a precooled heat sink until the compound solidifies.

7. Tear off the filter paper and allow the specimen within the cryostat to reach the same temperature while the microtome is being chilled.

8. Carefully trim the specimen block to a trapezoid shape using a prechilled razor blade.

Freeze Sectioning

1. Carefully mount the chuck containing the specimen in a microtome with the trapezoid surface parallel to the knife edge. Slowly retract the chuck until it clears the knife edge.

2. Set the section thickness at 8 to 10 mm in the microtome according to the manufacturer's instructions and prepare a smooth surface on the block by rapid speed cutting (e.g., one cut per second) until it is close to the specimen.

3. Before sectioning the sample, flip the plastic roll-plate down for collecting sections. The plate should just be touching the knife and lie parallel to the knife edge, but slightly behind the cutting edge of the knife.

4. Start sectioning by turning the crank. Once sections roll up at the cutting edge of the knife, lift the plastic roll plate and carefully expand the section onto the knife using a fine brush.

5. Transfer sections onto warm prelabeled and precoated glass slides by carefully overlaying the slide onto the section (Figure 12.2).

6. Repeat step 4 and step 5 until sectioning is complete.


Note: Frozen sections should be immediately fixed except for doing enzyme his-


FIGURE 12.2 Diagram of thin cryosectioning ribbons and mounting on a coated glass slide.

FIGURE 12.2 Diagram of thin cryosectioning ribbons and mounting on a coated glass slide.

1. Prepare a moisture chamber by adding a minimum volume of distilled water to an appropriately sized tray and placing a number of plastic or glass Pasteur pipettes on the bottom parallel to each other.

2. Place slides with sections on the pipettes in the moisture chamber and cover the sections with an appropriate volume of 4% PFA fixative or 10% buffered-formalin.

3. Cover the chamber and allow fixation to proceed for the desired time at room temperature.

Note: An optimal fixation time should be determined empirically for each tissue sample. In general, a range from 20 min to 8 h should be tested.

4. Carefully aspirate off the fixation solution and rinse with PBS for 6 min. Repeat washing with PBS three times.

5. Carry out dehydration by replacing PBS with a series of ethanol (40%, 50%, 75%, 95%, 100%, and 100%), 4 min each.

6. Air-dry the slides and store them in a slide box in an airtight container with desiccant at -80°C until use.

Note: Slides should be prewarmed to room temperature prior to use.


This section describes successful techniques for in situ hybridization using a radioactive probe. Procedures include dewaxing and rehydration/hydration of sections, protease digestion, DNase or RNase treatment, preparation of isotopic probes, hybridization and detection of cellular DNA or RNA.


Paraffin wax sections prepared in Part A must be subjected to complete dewaxing to allow treatments with protease, DNase, or RNase as well as penetration of the probe into the tissue. Dewaxing of sections can be completed readily with xylene. For unembedded cultured cells and cryosections, dewaxing is not necessary.

1. Transfer sections from the freezer to a slide rack and allow them to reach room temperature for a few minutes prior to use.

2. Fill three staining dishes with fresh xylene and two with 100% ethanol.

3. Dewax the sections by placing the slide rack into a series of three changes of xylene, prepared at step 2, for 3 min in each solution.

4. Wash the sections in two changes of 100% ethanol, prepared at step 2, for 3 min each.

5. Air-dry the slides and proceed to protease digestion.

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