Note

3.2.1.1. Dissection

1. Set up guillotine and surgery instruments for brain dissection.

2. Pour methyl butane into a stainless steel beaker and place in dry ice. Monitor temperature until it reaches -30 to -35°C. It is very important that temperature is kept within this range by placing the beaker back and forward in dry ice.

Fig. 3. (opposite page) Detection of Perl mRNA by in situ hybridization in the rat suprachiasmatic nucleus. (A) Film autoradiography of a coronal hypothalamic section of a rat sacrificed during the light phase of a light-dark cycle, and hybridized with 35S-labeled rPerl antisense riboprobe. (B) Fluorescent confocal photomicrograph of a coronal hypothalamic section of a rat sacrificed during the light phase of a light-dark

Fig. 3. (continued) cycle, and hybridized with digoxigenin-labeled rPerl antisense riboprobe, detected with Cy3 and visualized after excitation with 568-nm wavelength light. (C) The same section shown in (B) is double-labeled for arginine vasopressin by immunohistochemistry using PS 45 antibody (kindly provided by Dr. A. Gainer) and Alexa 488® goat anti-mouse secondary antibody (Molecular Probes), visualized after excitation with 488-nm wavelength light. Scale bar: 1 mm for A, 85 ^m for B and C.

3. Make a strainer with the Styrofoam cup by making holes at the base and cutting the wall so that only one third of the cup's wall is left. This strainer should fit comfortably in the stainless steel beaker.

4. Decapitate the animal. Depending on the mRNA species to label, decapitation without anesthesia may be recommended.

5. Carefully dissect the brain, taking special care to preserve the ventral surface, particularly at the optic chiasm region.

6. Place the brain on the base or the strainer, with the ventral side facing up. Brain should be symmetrically positioned on the strainer.

7. Dip the strainer in the methyl butane at -30 to -35°C and clamp the strainer so it stays in position for 3 to 5 min (see Note 23).

8. Place the brain in powdered dry ice. Leave the brain covered with dry ice for 5 min.

9. Remove the brain and shake all dry ice off its ventral surface. Put a drop or two of embedding matrix on your finger and rub it against the ventral surface of the brain. Embedding matrix should get in between all the grooves of the ventral surface of the brain. Return the brain immediately to dry ice and sprinkle powdered dry ice onto its ventral surface. Allow the embedding matrix to freeze and apply another coat of embedding matrix. There should be a 1- to 2-mm layer of embedding matrix on the ventral surface of the brain after two or three coats have been applied (see Note 24).

10. Wrap the brain in tape-labeled aluminum foil and transfer the brain from dry ice directly to the -80°C freezer until sectioning. Brains can be stored at -80°C for several months.

3.2.2. In Vitro Transcription of Radiolabeled Riboprobe

3.2.2.1. DNA Template

Templates are prepared as indicated in Subheading 3.1.2.1.

3.2.2.2. Riboprobe Synthesis

Radiolabeling of RNA probes is based on the tagging of specific RNA bases with a radioactive isotope, and the subsequent autoradiographic detection of the hybridized probe (see Note 25). In this protocol 35S-UTP is used as a source of uracils for the riboprobe. RNA probes can be also labeled with other radioisotopes such as 33P or 32P, provided that the adequate radiolabeled nucleotide is used.

1. Using 1 Mg of template DNA perform the labeling reaction according to the instructions of the MAXIscript Kit (Ambion), which contains all the necessary components with the exclusion of the radioactive nucleotide (see Note 16). Perform the optional DNase I treatment.

2. After DNase treatment, bring the reaction volume to 100 ^L with DEPC-treated water. Take 1 ^L of this final volume, dilute 1:100 in DEPC-treated water, and count 2 pL of this dilution in the scintillation counter to obtain your initial counts per minute (cpm).

3. Use a phenol/chloroform extraction followed by ethanol precipitation to purify your probe out of the remaining 99 pL of each reaction product. Resuspend each reaction product in 100 pL of 50 pM dithiothreitol (DTT) in DEPC-treated water, and repeat the counting procedure to obtain your final cpm. The percentage of incorporation, calculated using the initial and final cpm and volumes, should be higher than 60 to 70%. The final cpm will also be used to calculate the amount of probe per volume of hybridization buffer (see Subheading 3.2.3.3.). 35S-UTP-labeled riboprobes can be stored at -80°C for up to 2 mo. For other radioisotopes, this time will depend on the isotope half-life.

3.2.3. Tissue Sectioning, Prehybridization, and Hybridization

3.2.3.1. Tissue Sectioning

For this ISH protocol brain sections are mounted on microscope slides throughout the procedure. Before sectioning, microscope slides are coated with Vectabond™ according to the manufacturer's instructions. Brains are cut into 10- to 20-pm-thick sections in a cryostat and mounted immediately. Brains and sections are kept frozen all the time throughout the procedure. After cutting, the slides with sections are kept at -80°C in slice boxes within freezer bags with desiccant. Sections can be kept for several months at -80°C, although unnecessary long-term storage should be avoided. The integrity of the sections during cutting and mounting is critical for successful labeling of neuroanatomically defined areas.

3.2.3.2. Prehybridization

1. Remove slides from -80°C freezer and place them on slide racks.

2. Dip in 4% PFA for 5 min at room temperature.

4. Incubate 10 min in TEA-HCl/acetic anhydride.

5. Quickly rinse in 2X SSC.

6. Dehydrate in ethanol series prepared with DEPC-treated water as follows: 1 min in 70%, 1 min in 80%, 2 min in 95%, 1 min in 100%.

7. Delipidate by incubating in chloroform for 5 min (see Note 26).

8. Wash 1 min in 100% ethanol followed by 1 min in 95% ethanol.

9. Let air dry in a clean area. Slides can be left for a couple of hours while drying. While slides dry, set a high-humidity incubator to 37°C.

10. Mix equal amounts of 4X SSC and deionized formamide and pipet approx 50 pL of this solution on each slide. Cover slip each slide with clean glass cover slips so that the whole surface of the tissue section is embedded in the solution. Place slides in a capped plastic container and place the container in the 37°C incubator. Incubate 20 to 60 min.

3.2.3.3. Hybridization

1. Prepare the hybridization solution by diluting 108 cpm of probe per milliliter of hybridization buffer B with freshly added DTT.

2. Remove the plastic containers one at a time from the incubator. Remove the cover slips, carefully shake out any 50% 4X SSC/50% formamide left, and pipet approx 50 ^L of hybridization solution. Cover with a new glass cover slip (see Note 27).

3. Place a cap (i.e., from a conical tube) filled with water inside the plastic container and place the slides back into it. Alternatively, create a high-humidity chamber by placing on the bottom of the plastic container a piece of paper towel wetted with DEPC-treated water. Cover the plastic container, making sure it has a good seal. Otherwise, seal with Parafilm. It is extremely important that slides are in a high-humidity hermetic chamber; otherwise they will dry out.

4. Place the plastic container back into the incubator. Once every slide is covered with hybridization solution, and all plastic containers are back in the incubator, set the temperature of the incubator to 55°C and incubate overnight.

3.2.4. Post-Hybridization

1. Set an incubator or a water bath to 52°C and warm up enough 50% formamide/ 50% 2X SSC solution for two washes.

2. Set up three beakers with 1X SSC solution.

3. Remove the plastic containers from the incubator one at a time, remove each slide, and dip it in the first 1X SCC beaker. If the slides did not dry out after overnight hybridization, the cover slip should come loose easily. Rinse briefly in each of the two remaining 1X SSC beakers and incubate slides in slide racks in 1X SSC for 10 min. Transfer the slides to a second wash of 1X SSC for another 10 min. If many slides are processed, these two 10-min incubations can be done for a longer time. If hybridization was done with more than one probe, wash slides for each probe separately.

4. Incubate in 50% formamide/50% 2X SSC for 5 min at 52°C. For this incubation slides hybridized with different probes can be pooled together.

5. Transfer to fresh 50% formamide/50% 2X SSC and incubate for 20 min at 52°C.

6. While this incubation takes place, warm up an appropriate volume of RNase buffer in a 37°C incubator. This should be done in the RNase area of the laboratory.

7. Transfer materials to the RNase area and rinse two times in 2X SSC at room temperature for 1 min each (see Note 28).

8. Add RNase A to the RNase buffer to a final concentration of 100 mg/L and incubate the sections in RNase solution for 30 min at 37°C.

9. Remove the slide racks from the incubator and set it to 52°C.

10. While the incubator reaches 52°C, wash the slides two times in 2X SSC at room temperature for 5 min each, or longer if necessary.

11. Incubate in 50% formamide/50% 2X SSC for 5 min at 52°C.

12. Wash in ethanol series, diluted in 0.1X SSC instead of water, as follows: 3 min in 70%, 3 min in 80%, and 3 min in 95%.

13. Rinse the slides quickly in distilled water.

14. Wash in 70% ethanol diluted with distilled water, for 3 min.

15. Air-dry the slides and lay them out on autoradiographic cassettes.

16. In a dark room, place an autoradiographic film in each cassette, making sure that the emulsion side is against the sections. Cover the cassettes and place in a safe, dark area.

17. Develop the films after 2 d. Optimal exposure time must be determined empirically (see Note 29).

18. Autoradiographic images can be scanned and digitalized to estimate optical density. The optical density of the SCN can be normalized to the optical density of the surrounding hypothalamus (see Note 30).

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