Immunofluorescence - cell surface protein detection - How do I detect only cell surface expressed prot. (Jul/06/2005 )

Greetings

My first post!

I am a complete newbie when it comes to antibody work and immunofluorescence so all help appreciated.

The Story: We have a protein, expressed from a recombinant virus, that is natively expressed on the cell surface. We wish to prove that the protein is being cell surface expressed.

We have shown by immunofluorescence and acetone fixation that the protein is being expressed and we can detect it using polyclonal sera.

From my reading we need to fix the cells so as not to permeabilize the membrane to determine if the protein is being expressed on the cell surface. Am I correct and does anyone have a method I could thieve (possibly formaldehyde, gluteraldehyde, etc)?

Thanks for your time,

Scott.

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Hi DrScott,

Could to see another quality researcher with a great name. Anyway I haven't done much in the way of cell surfrace fluroescence but my fixation protocol is simply to grow cells on glass coverslip. fix with 4% Paraformaldehyde in PBS for 15-20 minutes. I also find that the aldehyde groups tend to autofluoresce so to help quench this I do a step in 150mM Glycine for 15-20minutes and then block in 5%BSA and add all antibodies in the presence of BSA. Normally I would add 0.3% Tx100 to permeabilise membrane but as long as your antibody recognises an extracellular epitope on your protein then you can leave this step off.

Hope this helps,

Scott

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What about doing a live cell stain? You'll need to work w/o detergent to keep the cell membrane intact. You'll need to use either culture medium or Dulbecco's PBS and work at 37C. But this will speed the antibody binding kinetics and reduce your inucbation times. Treat the cells with a blocking solution. You may not need to block since the cells are already grown in FBS. Add the primary for 1 hr. Wash gently with warm buffer w/o detergent. Then add a fluorescent secondary for 30 minutes. Wash with warm buffer. Then look at the cells. I've done this for a glycoprotein and for one other cell surface epitope. You can actually watch the staining evolve if you have a directly conjugated primary. As a control, you may be able to destroy the surface epitope with certain proteases--you'll need to check if there is a site on your protein. Alternatively, preincucbate your primary with 10 fold excess immunizing peptide for 1 hour at 37C prior to adding it to the cells.

I imagine you could do something similar with FACS analysis or even scrape the stained cells off the plate and sort live cells after cytochemistry.

As another idea, you may be able to biotinylate your surface epitope. Do an IP with your antibody or with SA-Agarose beads, and detect with SA-HRP or your antibody/followed by secondary, respectively.

Other points to keep in mind:

Phenol red has green autofluorescence.
Tissue culture plastic is not always ideal for fluorescent imaging.
Antibodies should be of great quality to avoid nonspecific interactions with other native epitopes.
Membrane proximal epitopes may not be as bioavailable for antibody binding.
Fc receptors may interfere with analysis.

Good luck!

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Sounds good! Along with the paraformaldehyde idea I think I have some things to try.

I'll be doing the work in glass chamber slides with media lacking phenol red so hopefully I'll be reducing some background there. One of the bigger problems (I think) will be the use of polyclonal sera as the primary antibody. I will pre-absorb to the cells before using in this experiment to try and reduce non-specific binding. It works beautifully on acetone fixed cells so lets hope.

Thanks for the replies.

Scott.

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