2) Pretrim the block and place it in the cryostat; for 20-30 minutes prior to cutting to equilibrate the block to the cryostat temperature. Cut 10-14 µm sections on the cryostat and transfer section ribbons onto slides (subbed slides not necessary). All the sections from one block may usually be fit on a single slide.
3) Warm the slides at 50-60°C for 2-3 minutes and then allow them to air dry for about 1 hour. In my experience, slides may be kept for several days sitting on a benchtop at this stage with no apparent loss of enzyme activity.
4) Fix the tissue sections in 2.0 % glutaraldehyde; in PBS for 15-20 minutes at room temperature in a coplin jar.
5) Wash the slides two or three times in PBS for 5 minutes/wash.
6) After the last wash, remove excess PBS with a tissue and place the slides on a level surface in a moist airtight chamber. A Tupperware box with water-soaked blotting paper works well for this purpose. To keep the slides off the paper, they should be placed on two lengths of plastic 5 ml or 10 ml pipettes which rest on top of the paper block.
7) Allow the remaining PBS on the slides to dry (5-10 min at room temp). The slides may be left to dry for a few hours at this point with no apparent loss of enzyme activity.
8) Apply 50 µl of a prewarmed staining solution + X-gal; to each slide and cover with a cover slip (50 µl suffices for a 22 x 40 mm cover slip - adjust accordingly for different sizes).
Staining solution: 50 ml total (store at room temperature
in a foil-wrapped 50 ml conical tube in the dark - stable for months).
1.8 ml of 0.2 M Na2HPO4
0.7 ml of 0.2 M NaH2PO4
1.5 ml of 5 M NaCl
50 µl of 1 M MgCl2
3 ml of 50 mM K3(Fe(CN)6)
3 ml of 50 mM K4(Fe(CN)6)
bring to 50 ml with ddH2O
X-gal 30X stock solution: store at -20°C
10% X-gal in N,N-dimethylformamide
Important: To avoid instantaneous precipitation of X-gal crystals; when applying solution to slides, it is necessary to prewarm these solutions. Generally, while the slides are in the fixative or washes, I aliquot out the required amount of staining solution into an eppendorf tube and incubate in a 42°C temp block for about 5 minutes. I then add 1/30 volume of the X-gal stock solution to this tube, quickly vortex or shake, and place it back in the 42°C temp block. I apply the staining solution + X-gal onto the dried slides as quickly as possible, put on all the cover slips, and immediately put the sealed chamber in a 37°C incubator.
9) Allow the staining reaction proceed for as long as necessary (from a few minutes to overnight).
10) Examine the slides periodically under a dissecting microscope. When the staining appears to be complete, float the cover slips off in PBS and wash the slides 2 x 5 minutes in PBS.
11) At this point, the slides may be dehydrated through a graded ethanol series and mounted in DPX; (Fluka) or other suitable mountant. The ethanol series also helps remove X-gal crystals deposited on the tissue sections during the staining reaction. Alternatively, the slides may be simply mounted in 70% glycerol in PBS after the final PBS wash (in my experience, the glycerol mounting; gives better morphology for photomicroscopy but is less suitable for long-term storage of slides).
12) For Hoechst; labeling; of nuclei, do not dehydrate the sections.
Instead, pipette 0.5-1.0 ml of a 1 µg/ml solution of Hoechst 33258
in PBS directly onto each slide. After 1-2 minutes, drain the dye solution
off, rewash the slides in PBS a few times (5 minutes each) and mount in 70%
glycerol/PBS.
1) Fix the tissue of interest in 2.0% formaldehyde in PBS at 4°C. For Drosophila heads, first remove the proboscis and fix the head capsule for 60-90 minutes. For other types of tissues, adjust the fixation time according to their relative size and estimated efficiency of infiltration.
2) Wash the tissue in PBS and transfer to 12% sucrose in PBS at 4°C. Allow the sucrose solution to infiltrate the tissue for 16 hrs.
3) Remove the tissue from the sucrose solution and submerge it in a drop of O.C.T. Tissue Tek; Compound (Miles Scientific). Allow the tissue to be permeated by the O.C.T. compound for 10-30 minutes at room temperature and then embed the tissue in frozen O.C.T. compound using an ethanol-dry ice bath (see preceding protocol.)
4) Place the frozen block containing the tissue to be sectioned in the cryostat ;chamber for at least 20 minutes to equilibrate it to the cryostat temperature (-14°C to -18°C).
5) Cut 10-14 µm sections on the cryostat and load onto freshly gelatinized slides.
6) Heat slides briefly at ~40°C on a drying plate for no more than one minute to dry the sections onto the slides. Overheating may adversely affect the antigenicity of the protein of interest and the morphology of the tissue.
7) Fix the sections immediately in 0.5% formaldehyde in PBS for 20-60 minutes at room temperature. Slides may be stored in this fixative at 4°C for a few days if necessary.
8) Wash the slides two or three times for 2-3 minutes/wash in PBS.
9) Block the slides for 30 minutes in PBSG (0.2% BSA, 1% goat serum,
0.01% saponin in PBS).
10) Wash the slides through several changes of PBS/0.01% saponin.
11) Apply 75-150 µl of the primary antibody at the appropriate dilution in PBSG; to each slide. Unless an excess of antibody solution is used, gently place cover slips on the slides to evenly spread the antibody solution over the tissue sections (100 µl of solution suffices for a 22 x 40 mm cover slip). Incubate the slides in the primary antibody for 30-60 min at room temperature or overnight at 4°C in a moist, airtight chamber (ie. a Tupperware box containing wet blotting paper).
12) Wash the slides through several changes of PBS/0.01% saponin
(float off the cover slips in the first wash).
13) Incubate the sections with the secondary antibody at the appropriate
dilution in PBSG exactly as described in step 11 for the primary antibody.
For Bio-Rad Laboratories HRP-conjugated goat-anti-mouse IgG;, a 1:200 dilution
for 30-60 minutes at room temperature works well.
14) For an HRP-conjugated secondary antibody, the slides are again washed through several changes of PBS/0.01% saponin and then incubated in 0.5-1.0 ml of staining solution per slide without cover slips or 100-150 µl per slide with cover slips. The staining solution consists of: 0.5 mg/ml diaminobenzidine (DAB), 0.003% H2O2, 1.5 mM CoCl2, and 1.5 mM NiCl2. The CoCl2 and NiCl2 are optional components; in their absence, a brown product forms, whereas in their presence, a more intense blue-black product is obtained. Since the extent and localization of background staining can vary considerably between intensified and non-intensified preparations, it is often worth trying both methods in parallel.
NOTE: DAB; is thought to be a potent carcinogen;
and is to be handled with care. All DAB-containing tubes, pipette tips,
used DAB solutions and similar materials should be immersed overnight in 100%
bleach and disposed of according to the proper regulations.
15) Monitor the staining reaction under a dissecting microscope.
When the reaction has proceeded to completion (usually 5-30 minutes), it
is stopped by washing the slides through a few changes of PBS.
16) For compound microscope viewing and photomicroscopy, the sections may either be dehydrated through a graded ethanol series and mounted under cover slips in a stable mountant such as DPX; (Fluka Biochemie), or they may be mounted under cover slips in 70% glycerol/PBS after air-drying without dehydration. In general, the glycerol mounting; is superior for photomicroscopy but is not suitable for stable, long-term storage of samples.
17) For Hoechst; flourescent counterstaining of nuclei, do not dehydrate the sections after step 15. Instead, pipette 0.5-1.0 ml of a 1.0 µg/ml solution of Hoechst 33258 in PBS directly onto each slide, allow to sit for 1-2 minutes, then rinse off through a few changes of PBS and mount in 70% glycerol/PBS as described above.