Appendix D: Rapid Protocols
D.1 HEMATOXYLIN & EOSIN STAINING
D.1.1 Start Notes
The following protocol is to be performed at room temperature but hematoxylins can
be prewarmed to 50 60 C to increase the signal (usually this is not necessary and is not
recommended if using fresh dyes).
Ideally, all material should be rinsed with distilled water and allowed to dry before the
process, and reagents should be fresh. However, they can be used for several series of
slides without consequence. Waters should always be replaced, though. Absolute ethanol
just before the final clearing step should be replaced more often, as it tends to get
hydrated and produce a whitish suspension when sections are transferred to xylenes. If
this happens, wash the sections thoroughly with fresh xylenes.
D.1.2 Procedure
1. Fixation
Bouin’s fixative 36 48 h
Wash in good quality tap water 3 3 1 h
2. Dehydration
70% ethanol 1 3 30 min
95%/96% ethanol 2 3 15 min
Absolute ethanol 3 3 30 min
3. Intermediate infiltration
Xylenes 3 3 15 min
4. Infiltration
Molten paraffin At least o/n
5. Cast, section, and collect onto coated slides
6. Deparaffination
Xylenes 2 3 30 s
7. Rehydration
Absolute ethanol 2 3 30 s
95%/96% ethanol 2 3 30 s
70% ethanol 1 3 30 s
Brief rinse in distilled water
Distilled water 1 3 6 min
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APPENDIX D: RAPID PROTOCOLS
8. Staining
Harris’s hematoxylin 2 min
Differentiate in tap water 2 3 2 min
Brief rinse in distilled water
70% ethanol 1 3 3 min
Alcoholic Eosin Y 1 min
9. Dehydration
70% ethanol 1 3 30 s
95%/96% ethanol 2 3 30 s
Absolute ethanol 2 3 30 s
10. Clearing
Xylenes 1 3 30 s
Drying At least 1 h under a fume hood (o/n on benchtop)
11. Mount with DPX or similar resin and allow drying at least o/n under a fume hood.
D.2 RAPID PROTOCOL FOR FLUORESCENCE
IMMUNOHISTOCHEMISTRY
D.2.1 Start Notes
The following protocol is to be performed at room temperature unless specifically
stated otherwise. Neutral-buffered formalin is recommended for most cases and sample
sizes. For small samples, paraformaldehyde can be used, with better results. Other fixatives should be effective, like Davidson’s (which provides very good results for IF at least),
but avoid Bouin’s and Zenker’s.
Include positive controls whenever available but always add negative controls to the
process, at least one per experimental treatment or series of slides. Ideally, each slide
should be accompanied by its respective negative control. As such, it is recommended that
two slides are prepared from each specimen. The slides treated as negative controls will
not receive the primary antibody, being treated with buffer only. All other steps of the
procedure should be the same as described below.
All incubation steps are to be performed in a covered humidified chamber for IHC,
with distilled water over the bottom. Keep the tray of the chamber, where the slides are
placed, clean and dry to avoid loss of liquids by capillarity.
Rinsing is conveniently done using a washbottle with sterilized PBS.
Washing can be done in Coplin or Hellendahl jars containing sterilized PBS. The change of
PBS in these jars between steps is highly recommended. Avoid washing the negative controls
in the same jar as the other slides right after the incubation with the primary antibody.
The use of Dulbecco’s PBS is preferred. Use the same PBS for all steps and solutions.
Prepare the antibody solutions using the blocking buffer as diluent (2% BSA in PBS
with 0.1% Triton X-100).
Do not reuse coverslips.
The following protocol is conceived for secondary antibodies prelabeled with a fluorochrome (red, like Cy3, is recommended for most cases), being thus peroxidase enzyme-
APPENDIX D: RAPID PROTOCOLS
253
independent. Slides should be kept in the dark as much as possible after the secondary
antibody incubation.
D.2.2 Procedure
1. Fixation
Neutral-buffered formalin 24 h
Wash in good quality tap water 3 3 1 h
2. Dehydration
70% ethanol 1 3 30 min
95%/96% ethanol 2 3 15 min
Absolute ethanol 3 3 30 min
3. Infiltration
Xylenes 3 3 15 min
4. Infiltration
Molten paraffin At least o/n
5. Cast, section, and collect onto precoated slides (e.g., Polysine)
6. Deparaffination
Xylenes 2 3 30 s
7. Rehydration
Absolute ethanol 2 3 30 s
95%/96% ethanol 2 3 30 s
70% ethanol 1 3 30 s
Brief rinse in distilled water
PBS 1 3 6 min
8. Permeabilization/antigen retrieval
0.1% Triton X-100 in PBS for 15 min
OR
0.05% Trypsin, pH 7.8, with calcium chloride (0.1%) for 15 min at 37 C
Fast rinse in PBS (1 3 )
Wash in PBS (2 3 30 s)
9. Blocking
Add 200 µL of blocking solution to the slide: 2% BSA in PBS with 0.1% Triton X-100
Place coverslip
Incubate for 30 min
Remove coverslip
Fast rinse in PBS (1 3 )
Wash in PBS (2 3 30 s)
10. Primary antibody
Add 200 µL with the working primary antibody solution to the slide, diluted to 10 µg/L if no optimal concentration is specified.
Place coverslip
Incubate the slides o/n at 4 C
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APPENDIX D: RAPID PROTOCOLS
Remove coverslip
Fast rinse in PBS (1 3 )
Wash in PBS (2 3 30 s)
11. Fluorochrome-labeled secondary antibody
Add 200 µL with the working labeled secondary antibody solution to the slide,
diluted to 10 µg/L if no optimal concentration is specified.
Place coverslip
Incubate the slides for 2 h, in the dark
Remove coverslip
Fast rinse in PBS (1 3 )
Wash in PBS (2 3 30 s)
12. Mounting
Place four drops ( 200 µL) of aqueous mounting agent with DAPI
Analyze immediately (keep preparations in the dark)
D.3 BASIC PREPARATION OF SAMPLES FOR TEM
D.3.1 Start Notes
The following protocol is to be performed at room temperature unless stated otherwise.
Cacodylate buffer (working solution) is 0.1 M, pH 7.4.
Glutaraldehyde (2.5% m/v) and osmium tetroxide (1% m/v) solutions are prepared in
cacodylate buffer.
Osmium fixative contains 0.1% m/v osmium tetroxide in 0.1 M, pH 7.4, cacodylate
buffer.
Epon refers to the final mixture according to Luft’s recipe (see Section 3.3).
Use flat rubber embedding molds for TEM, rinsed with acetone and allowed to dry.
Molds with a cavity size of 3 6 mm are adequate for most occasions but check compatibility with ultramicrotome specimen holder.
Samples processed until dehydration in 100% acetone can be further processed for
SEM.
D.3.2 Procedure
1. Fixation
Collect a fresh 1 2 mm specimen from biological sample. Dissect and handle in
fixative.
Glutaraldehyde fixative 1 2 h
Wash in cacodylate buffer 3 3 15 min
2. Postfixation
Osmium tetroxide o/n
Wash in ultrapure water 3 3 15 min
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255
3. Dehydration
30% acetone 10 min
50% acetone 10 min
70% acetone 10 min (can be used for archiving)
90% acetone 10 min
100% acetone 3 3 10 min
4. Intermediate infiltration
2:1 propylene oxide:Epon 30 min
1:1 propylene oxide:Epon 30 min
1:2 propylene oxide:Epon 30 min
5. Infiltration
Epon (low-pressure vacuum) 30 min
6. Casting
Samples 1 Epon in molds o/n (45 60 C)
Final hardening 1 2 h (70 C)
Air-cure decasted specimens At least o/n
7. Sectioning
Trim the block to a 1-mm-wide trapeze under a stereoscope using a thin, sharp,
blade
Section with the microtome at 200 400 nm thick (semithin sections)
Collect sections with a copper-wire scoop and blot them onto a microscopy glass
slide
Dry the slide on a hot plate ( 70 C) for about half an hour or more
Stain with a few drops of Toluidine Blue (sufficient to cover all sections) over the hot
plate until the edges of the dye begin to dry, wash in a gentle stream of running tap
water until all the excess dye is washed off, and return the slide to the hot plate.
Once dry, mount with an aqueous or resinous medium and check condition of the
specimen under the microscope
If satisfactory, section at 100 nm thick to obtain thin sections for TEM
8. Collect sections onto grids (e.g., Cu, 300 mesh), degreased with chloroform, and blot to
dry
9. Staining
Uranyl acetate 45 min
Rinse with ultrapure water
Blot to dry
Lead citrate (Reynolds) 5 min
Rinse with ultrapure water
Blot to dry