Hi! I´d like to ask for a protocol to detect GFP fluorescence after fixation and permeabilization.
This is the situation:
I´m using a MSCV-MLL-AF9-IRES-GFP plasmid to induce acute myeloid leukemia into mice in vivo. To do so, I infect mouse hematopoietic stem cells (HSCs) with a retrovirus containing the plasmid and then I transplant them into mice. After 10 weeks, the mice get sick (as I can see GFP fluorescence in blood).
When I analyze leukemia cells in vivo, I can see a nice GFP signal. However, after fixation and permeabilization, the GFP fluorescence is lost. I tried to detect it with an anti-GFP antibody, but it doesn´t work. I´ve done this many times changing some parameters (fixation and permeabilization reagents, incubation times, etc.) but nothing seems to work. This is the detailed protocol of what I do, in case you identify a potential mistake:
- Isolation of bone marrow cells from mice and resuspension in staining media (HBSS + BSA)
- Fixation with 1% PFA for 15´ on ice in the dark
- Permeabilization with permeabilization buffer (containing saponin) for 5´ at room temp in the dark
- Incubation with the GFP antibody for 1 h
- Wash with permeabilization buffer x2
- Resuspension of stained cells in PBS+DAPI
- Analysis by flow cytometry
When I run the samples in the flow, I can see cells have been properly permeabilized since most of them are DAPI+. However, I don´t see any GFP signal (I use AF488 anti-GFP antibody).
Please some recommendations on how to solve this!!
What is the pH of your paraformaldehyde? It should be around 7.4 for best preservation of GFP signal. If the pH is too basic, it will quench the GFP signal.
In another thread, Brian Lin at Massachusetts General Hospital suggested the following fixative (1% PLP - paraformaldehyde, lysine and periodate):
"Briefly, to make 100 mls, dissolve 1 g PFA in 60 ml of 60C dH2O. Solubilize with a few drops of 1 N NaOH on low heat and stirring.
Weigh out solid .1M Phosphates: 1.014 g Na2HPO4 (dibasic) and .392g NaH2PO4 (monobasic) and add it to the 1% PFA with stirring.
Simple paper filter it, and add to it 1.37 g of lysine mono-HCL, predissolved in 10 ml H20.
Finally, add in .21g Na Periodate and QC to 100 ml.
(like I said, this is very laborious, but it is an excellent fixative for people who are doing careful IHC, DAB or fluoresence in tissue. Also, the lysine and periodate appear to do some kind of complexing reaction, so the fixative needs to be made fresh, at most 6 or 7 hours before use."
Hope that helps some. Good luck!
I agree that the fixative is a good place to start troubleshooting.
Another suggestion would be 100% methanol, it can be a good alternative for staining cells with antibodies that don't work with PFA, and also permeabilizes the cells.
not in my experience, Matthias.
we routinely use methanol treatments for cells and zebrafish expressing a variety of transgenes. however, we usually use it for IF colocalization, not flow.
so methanol may reduce the GFP, but i really doubt it would kill it - especially if the GFP being detected is coming from an antibody that's applied post-fix
Article Effect of fixation procedures on the fluorescence lifetimes ...
Thanks for all the suggestions!
I´ve read other people´s concerns about using PFA that they were able to solve the problem by using 1% PFA instead of 4%. I always use 1% and I still loose my GFP signal.
I checked the pH of my PFA and it was neutral, so I discarded the pH variable.
Also, I don´t understand why I can´t detect GFP when I use an anti-GFP antibody (I´ve read a lot of comments of people using it satisfactory). I know it´s not a problem of permeabilization since I stain for other intracellular markers and I can see these very nicely.
Since your gfp is tagged to any cellular structure proteins, GFP will lost after cell permeabilization if GFP is not fully crosslinked by fixation. We use 4% buffered PFA and fix the cells for at least 20min, we can keep 50% of GFP in term of intensity. Hope this helps.
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