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Intracellular stain of nuclear receptor - (Jul/16/2009 )

Hi,

I'm trying to do intracellular staining of mouse dendritic cells. Since I only have non-conjugated antibody, I have to do indirect staining by secondary antibody conjugated with PE.

I've tried several fix and perm condition (90% MeOH, 70% EtOH, 0.1% Saponin at different time point and temperature), there is still no staining using Histone (H4) as my positive control. I've tried also eBioscience Foxp3 staining reagent too, the result is the same.

I'm not sure is it the problem of perm condition is not optimized or the problem of indirect stain. Is anyone had experience before or can give me some advice in doing indirect staining of intracellular antigen? For antibody concentration, I'm using the highest dilution recommended by the manufacturer (for primary, about 1:50, seconday. about 1ug/1million cells).

Thank you very much for your kind advice.

-SMN-

Hi SMN,
Have you also tried triton-x for perming the cells?

Do you know if your histone antibody works with intracellular staining?

Are you fixing your cells before you perm them?

Have you tried DNAse treating your cells? This may help your antibody "get to" the histone epitope.

The BD BrdU kit had an alternative protocol for nuclear permeabilization. You can get it off the protocol pdf. I think (if my memory serves), the cells are fixed in 4% PFA, washed, frozen in 90% FBS/10% DMSO for at least 2 hours at -80 degrees, refixed with 4% PFA, and then washed with 0.1% saponin buffer.

Are you using ice-cold methanol for your perm steps?


Good luck and I hope this helps!

-miBunny-

Hi miBunny,

I didn't try triton-x but i tried 70% ice-cold Ethanol for 30 mins to perm the cells.

I think the problem of histone maybe as u suggested, besides H4. Since I want to avoid the DNAse treatment, can u suggest alternative control?

Thanks!!

-SMN-

thats a tough one. maybe restriction digests???

-miBunny-

For histone (p-H2A.X) labelling in cancer cells, I use the following protocol. It works very well. Maybe it would be helpful for your staining, too:

Reagents:
- 2% Paraformaldehyde
- Blocking buffer: 3% BSA, 0.05% Saponin (or Tween20 or Triton X-100) in PBS
- Rabbit primary antibody
- Secondary antibody

1. harvest cells, wash 1x with PBS
2. fix in ~100 ul PBS and 200 ul 2% paraformaldehyde solution while mixing (you can use any final concentration between 1-4% PFA)
3. leave the cells for 15 min at room temperature (not longer to prevent excessive autofluorescence!)
4. stop fixation by adding 1 ml cold PBS
5. spin off paraformaldehyde
6. add 0.1% Triton X-100 (0.5 1ml for 10^6 cells) and leave them for 10-30 min at RT
7. wash twice with blocking buffer
8. add primary antibody (1 2 ul)
9. incubate tubes 1h at RT
10. wash cells ONCE with wash buffer
11. add secondary antibody (1 ul) to cell pellet for 30-60 min
12. wash once
13. add RNAse and PI, if desired

Paraformaldehyde can be either bought as stock solution (methanol-free!) or prepared from powder.

To prepare from powder:
 weigh 10.0 g PFA in fume hood
 dissolve in 450 ml distilled water by warming up in the microwave shortly
 add few drops of 5M NaOH until the cloudy suspension will turn clear
 add 50 ml 10x PBS and adjust pH to 7.3
 aliquot into tubes and freeze, protect from light


Good luck!

-illuminated-

Thank you very much for your information.

All the best in your research too!

-SMN-

What I forgot to mention: ethanol fixation does permeabilise your cell, but it permeabilises it "too much". If the histone is a relatively small molecule, it will escape from the cell and you won't be able to detect it. In case of "my" histone this is the case, so paraformaldehyde fixation is crucial! I assume that histones are all of approx. the same size, so this might explain your results.

-illuminated-