Dear Richard,

Are you using PFA or methanol fixation? Methanol generally gives much less cytosolic staining since it doesn't fix soluble components very well - it is, however, very good for centrosome staining.

For pre-extraction I would use a microtubule stabilizing buffer (low salt, low pH) with saponin or Triton X100 (0.1-0.5%). PBS will destabilize microtubules and since microtubules help organize mitochondria it may interfere with your analysis.

A common microtubule stabilizing buffer is, for example, PHEM: 60 mM Pipes, 25 mM Hepes, 5 mM EGTA, 1 mM MgCl, pH 6.9

Detergent concentration and time depend on the cell type.

Hope that helps!

For immunostaining of live cells I've incubated cells for 15-20 min on ice using 0.05% saponin/0.5% BSA in PBS containing the appropriate primary antibodies, washed cells 2-3X with ice-cold PBS, fixed with 4% PFA and labelled with secondary antibodies in the presence of 0.1% saponin/0.5% BSA in PBS. Perhaps incubating with primary antibody for a longer period of time and staining on ice would improve the consistency of results while lessening the effects on organelle morphology. 

Respected Sir,

I have used 0.5% triton-X in 4% PFA for 5 minutes followed by fixation in 4%PFA  for 20 min. The time and concentration of triton X can be standardised according to proteins.

I would suggest Mitchison lab protocol, however, they are expert of microtubules and similar things, but not organelles. They use/suggest  high triton-x-100 (about 0,5% in buffer) for soluble actin extraction, for 30 seconds, and room temperature (?), next promptly fix the sample. 

Dear Richard,

Are you using PFA or methanol fixation? Methanol generally gives much less cytosolic staining since it doesn't fix soluble components very well - it is, however, very good for centrosome staining.

For pre-extraction I would use a microtubule stabilizing buffer (low salt, low pH) with saponin or Triton X100 (0.1-0.5%). PBS will destabilize microtubules and since microtubules help organize mitochondria it may interfere with your analysis.

A common microtubule stabilizing buffer is, for example, PHEM: 60 mM Pipes, 25 mM Hepes, 5 mM EGTA, 1 mM MgCl, pH 6.9

Detergent concentration and time depend on the cell type.

Hope that helps!

Jens,

Thank you for the response. I agree re cytosolic and centrosome staining and you make an excellent point about the potential value in switching from PBS to PHEM. We will try this straight away!

Rick

We had good results with the permeabilization protocol from Mary Dasso’s lab (Joseph et al., J. Cell Biol. 2002, 156:595-602). Cells are permeabilized for 3-4 min at 22ºC in 0.005% digitonin in “transport buffer”: 110 mM potassium acetate, 20 mM HEPES (pH 7.3), 2 mM magnesium acetate, 0.5 mM EGTA, 2mM DTT, + protease inhibitors (filter the solution before use) followed by paraformaldehyde fixation. The digitonin stock solution is at 50 mg/ml (5%) in DMSO, stored at –20ºC. We found this protocol to be particularly useful to lower the IF signal from soluble cytosolic proteins, e.g. in protein overexpression experiments.

I'd like to hijack this question if you all don't mind to ask a question of my own.  After permeabilization, what percentage of proteins diffuse out of the cell?  I'm curious about the degree to which proteins are bound to cytoskeletal features as well as other organelles. Do you all see a tremendous loss of cytoplasmic staining after pre-fixation permeabilization?  Cheers!

I would suggest that the answer to Michael Davis' question will be very protein and possibly even cell type dependent. Also, protein level. Specifically it is when you over-express a protein that it is much less likely to have binding partners that impart specific localization and you see mis-localization to membranes that are not "biologically relevant" or excessive staining in cytosol for soluble proteins.