Katherine A. Staskus
[This is a preliminary document!]
Tissue sections, 8 µM thick, are cut from paraffin blocks and are attached to slides (which have been coated with Denhardt's medium and acetylated) with a solution of 3% (v/v) Elmer's white glue in deionized distilled water (ddH2O). The slides are air dried and stored at room temperature.
[Note: tissues are fixed for 6-8 hr in fresh 4% (w/v) paraformaldehyde in phosphate-buffered saline (PBS) or 10% buffered-formalin]
KS: I have also successfully used organosilane-treated slides as a replacement for the Denhardt-coated slides. In this case the sections are floated onto the slide from a water bath and dried.
Denhardt treatment of slides (thickness: blue > green > gold > silver)
Wear gloves, rinse off powder to avoid talc artifact. touch only the edges of the slides!
Place slides in racks (50 per)
Soak slides in 1 M HCl for 30 min. (86.21 ml/liter)
Rinse in H2O, couple of changes.
Soak in absolute ethanol for 30 min., air dry.
3X (to 7X) denhardt's medium in 3X SSC, 65°C overnight
Make 50X denhardt's media as in Maniatis, p.448.
Use BSA, 99%, mostly immunoglobulins, NOT fatty acid free)
BSA Sigma A-7906
Ficoll Sigma F-9378
PVP Sigma PVP-360
1 g each / 100 ml
Dip very quickly once in water
Ethanol:acetic acid (3:1) for 20 min.
Air dry
Acetylate
Air dry
Silanization of slides
Wash slides in 2 N HCl for 5 minutes.
Rinse in distilled water for 1 minute.
Rinse in high grade acetone for 1 minute and air dry.
Soak slides in a 2% solution of organosilane in high grade acetone for 1 minute with
gentle agitation.
3-aminopropyltriethoxy silane, Sigma A-3648
Rinse slides in high grade acetone for 1 minute and air dry.
Store boxed at room temperature.
[from PCR in situ hybridization: protocols and applications
G.J. Nuovo, (1992) Raven Press, N.Y.]
Deparaffinization: The slides are placed in a rack in a 60°C oven for 60 minutes.
Two 5 minute washes in xylenes, with continuous agitation.
One 5 minute wash in absolute ethanol.
Air dry.
In situ PCR
pretreatments: none (or proteinase, experiment with conditions)
primer set: same primers as described in Embretson, et al. publications:
PNAS 90, 357 (1993) and Nature 362, 359 (1993)
reaction mixture: 10 mM Tris-HCl, pH8.3
50 mM KCl
0.01% (w/v) gelatin
1.5 mM MgCl2
200 µM dNTPs, 10 mM stock solution
1 µM each primer, 10 µM stock solution
slide setup/sealing process:
The reaction mixture is assembled and heated for 10 minutes at 94°C in a heating block.
Amplitaq DNA polymerase (PECetus) is added to a concentration of 0.15 U/µl (3% v/v) and the mixture is returned to 94°C. Aliquots are removed and pipetted onto dry deparaffinized tissue sections which are then overlaid with siliconized glass coverslips. (~5 µl for 18 x 18 mm coverslips, 10+ µl for 20 x 30 mm coverslips). The slides are placed in heat-sealable plastic bags (4 in. x 6 in., 2 mm thick); 2 slides/bag. 4-5 ml of mineral oil is added to each bag, air is removed, and the bags are heat-sealed and placed in the thermal-cycling oven.
post-PCR processing:
Remove slides from bags.
2 minute wash in chloroform, twice. Air dry.
Remove coverslips and dip slides briefly in fresh chloroform. Air dry for 5 minutes.
PBS for 5 minutes.
Dehydrate (5 minutes each 70%, 80%, absolute ethanol) and air dry.
In situ hybridization
pretreatments:
0.2 N HCl for 30 minutes
ddH2O rinse
0.15 M Triethanolamine (TEA) [1 M stock solution], pH7.4 for 15 minutes
ddH2O rinse
digitonin solution for 5 minutes:
0.05% digitonin (stock is 1% [w/v] in ethanol, stored at 4°C)
0.125 M sucrose (1.25 M stock solution)
60 mM KCl (1 or 2 M stock solution)
3 mM Hepes, pH7.4 (0.5 M stock solution)
ddH2O rinse
proteinase K for 15 minutes at 37°C:
5 µg/ml proteinase K (20 mg/ml stock solution in ddH2O, -20°C)
20 mM Tris-HCl, pH7.4 (1 M stock solution)
2 mM CaCl2 (1 M stock solution)
ddH2O for 5 minutes, twice
dehydrate and air dry
For RNA hybridizations, at this point acetylate the slides and set up the hybridization. For DNA, continue with the following pretreatments.
Add RNase mixture (enough to wet tissue) and coverslip:
100 µg/ml RNase A
10 U/ml RNase T1
2X SSC
incubate at 37°C in humidified chamber for 30 minutes
removed coverslips and wash slides in 2X SSC, twice for 5 minutes with constant
agitation.
post-fix for 2 hours in fresh 4% (w/v) paraformaldehyde in PBS
wash in 2X SSC for 5 minutes, twice
ddH2O rinse
acetylate:
place slides in 0.1 M TEA, pH7.4
add acetic anhydride, drop-wise, to a concentration of 0.25% while mixing
agitate slides for 10 minutes
ddH2O rinse
denature in 95% formamide in 0.1X SSC at 65°C for 15 minutes
ddH2O rinse
0.1X SSC at 4°C for 2 minutes
dehydrate
hybridization mixture: 10% (v/v) dextran sulfate
50% formamide
20 mM Hepes, pH7.4
1 mM EDTA
1X Denhardt's medium
1 mg/ml polyA
0.6 M NaCl
100 µM aurintricarboxylic acid (ATA)
110-140 µg/ml yeast RNA
100 mM dithiothreitol
1-6 x 105 cpm 35S-labeled probe/5 µl
stock solutions: 20% (w/v) dextran sulfate in deionized formamide, -20°C
5 M NaCl
10 mM ATA
free acid form is dissolved in water and adjusted to pH7.6
with 1 N NaOH, filtered and stored at 23°C in the dark.
2 M DTT made FRESH!
10X stock containing the Hepes, EDTA, Denhardt's and polyA
The Denhardt's is made with nuclease-free BSA
This solution is not filtered, rather, it is spun for 10 minutes
in a clinical centrifuge and aliquots are stored at -20°C.
Required amount of probe is ethanol precipitated in the presence of ~30 µg yeast RNA with 0.3 M NaAcetate and 2 volumes of absolute ethanol. Probe is pelleted in the microfuge, washed with 70% ethanol and dried in the speedvac. Probe is resuspended in the volume of ddH2O required for the hybridization mixture, boiled 5 minutes, quenched in an ice slurry and spun briefly in the microfuge. The remaining components of the hybridization mixture are added.
Hybridization mixture is pipetted onto the tissue section which is then overlaid with a siliconized coverslip and sealed with 2 layers of rubber cement around the edges. Hybridization is carried out for 2-3 days at 42°C.
post-hybridization wash:
Rubber cement is peeled off of the slides and the coverslips are removed under 2X SSC. Slides are incubated in 2X SSC at 55°C for 1 hour and then washed for 2-3 days in formamide-containing wash buffer: 50% formamide
20 mM Hepes, pH7.4
0.6 M NaCl
5 mM DTT
The slides are placed in 2X SSC for 5 minutes and then dehydrated through graded alcohols (70%, 80%, absolute) containing 0.3 M NH4Acetate and air dried.
development:
The slides are dipped in photographic emulsion at 43°C (Kodak NTB-2 which has been diluted 1:1 with 0.6 M NH4Acetate) and air dried for 2 hours. Slides are sealed in light-proof boxes with desiccant and placed at 4°C for period of exposure. Slides are developed in Kodak D-19 developer and Kodak fixer and counterstained with Hematoxylin or Wright's stain.
Photographic emulsion (Kodak NTB-2) is stored in a light-proof container in a radioisotope-free 4°C environment. Working emulsion is made by diluting new emulsion 1:1 with 0.6 M NH4Acetate: Remove emulsion from the cold and let it warm to room temperature in its light-proof container for 1-1 1/2 hours. In the darkroom (without safelight) place the bottle of emulsion and an equal volume of 0.6 M NH4Acetate (in a bottle with cap - which can accomodate the double volume) into a 43°C waterbath - for one hour to melt the emulsion and equilibrate temperature. Gently pour the emulsion into the NH4Acetate. To do this tilt the bottle of NH4Acetate and pour the emulsion into it down the side of the container. You want to avoid the formation of air bubbles which can bedifficult to remove once they are there. Tightly cap the bottle, tip it on its side and slowly roll the bottle in your hands for a few minutes to mix the contents. Pour the diluted emulsion into 2 containers and store at 4°C as previously described. Diluted emulsion is good for 3-4 months.
Coating slides with emulsion:
Remove diluted emulsion from 4°C and warm at room temperature for 1 hour. In the darkroom, remove the emulsion from the light-proof container and place in a 43°C waterbath for 45 - 60 minutes. Dip a blank slide, walk out of the darkroom and look at the coating on that slide. If the slide is covered with bubbles this means you have many bubbles in the emulsion. You can remove many of them by dipping blank slides into the emulsion and discarding them. If the coating looks clear and watery (it should be smooth and opaque) the NH4Acetate has separated out. In this case, either gently stir the emulsion with a blank slide, or cap the bottle and roll to mix as described before. After dipping, place the slides on end in a rack (with paper toweling underneath to absorb what solution drains from the slides) and let dry in the darkroom for 1-2 hours. Place the slides in a light-proof slide box which contains dessicant and store at 4°C for period of exposure. We wrap our slide boxes in a double layer of aluminum foil to ensure that light will not enter.
Development of slides:
Remove slide boxes from 4°C and warm for 1 hour at room temperature. In the dark room, remove the slides from the boxes and place in a glass rack which fits the glass slide-staining jars. Develop the slides in Kodak D-19 developer (made according to manufacturer) at room temperature for 3 minutes. DO NOT aggitate slides - as this may rip the swollen emulsion off of the slides. Dip the rack of slides once in room temperature water. Fix the slides in Kodak Fixer (made according to manufacturer) for 3 minutes. Rinse the slides for 1 minute in water, twice, and air dry. Do not turn on the lights until the slides are out of the fixer. Stain slides with appropriate stain (H&E, Wright's, etc.), dry, clear in xylenes and mount under coverslip with permanent mounting media.