Methods and Techniques Discussion > Immunofluorescence (IF)

Thick tissue sections



For a new project, I will have to stain thick sections (at least 100 μm) and analyze them by confocal microscopy.
We have non-fixed frozen tissue, formol-sucrose fixed frozen tissue and paraffin blocks.

I already tried paraffin sections of 100 μm on a glass slide, but they fell of in the first toluene bath.
Now I will try with frozen sections (fixed and non-fixed) on a glass slide.

But I was wondering if it would be better to use floating sections? And on what type of tissue, frozen or paraffin?
However, I have no experience with this technique.
I have a lot of questions like:
How do you put the tissue in a well without breaking it? And how do you place it on a glass slide afterwards?
Which dilution and how long should I use Triton X-100 to make the tissue permeable?
And do I need do use an optical-clearing solution like BABB (benzyl-alcohol - benzyl-benzoate) before using confocal microscopy on these thick sections?

Does anyone have a immunofluorescent staining protocol for these thick (floating?) sections?
(I normally only stain thin sections)

Thanks a lot!


Hey, I know this is an old post but just in case you are still interested, I do wholemount staining with skin and adipose tissue, and it works well with the following protocol.

I think it's best to use non-fixed, and just have a very light fixing step with Acetone, that way you can skip antigen retrieval steps. What I do is get a 96 well plate, throw the tissue in PBS at the top, and fill all the wells with the solutions you need for each step. This is actually much easier than working with slides, no more blotting away solutions, you just pick the tissue out with tweezers and move it down a row.

1. Dissect tissue and finely mince in 1x PBS (it's easy to cut thin slices if the tissue is frozen, you work on ice and use a razor or microtome blade).

2. Fix tissue for 30 min, RT on rotating platform (-20C Acetone)

3. Wash 3 times, first wash with 50 mM Glycine 5 min, then 2x 10 min in PBS (Glycine to reduce bg).

4. Permeabilize cells and block non-specific binding in 1 mL B1 buffer at RT on rotating platform (1 hour)

5. Incubate tissue with appropriate dilution of primary antibody in B1 buffer at 4 ˚C on shaking incubator o/n

6. Wash 3 times with 1 mL B2 buffer, 10 min on rotating platform

7. Incubate with appropriate dilution of secondary antibody in B1 buffer and DAPI/DRAQ5/ToPro/Bodipy...etc. RT on shaking incubator, 900 rpm (o/n)

8. Wash 3 times with 1 mL B2 buffer, 10 min on rotating platform

9. Mount specimen in 80% glycerol/1x PBS

B1 buffer    1% BSA, 5% normal goat serum, 0.3% Tween PBST in PBS, 0.45 um filter sterilize, keep at 4˚C

B2 buffer    0.1% Tween PBST in PBS

PBST (phosphate buffered saline)   3% Tween 20


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