Home >
Community >
What protocol can I use for the pre-fixation permeabilization of...
Upvote
27
Downvote
+ Molecular biological techniques
+ Saponins
+ Proteins
+ Immunofluorescence
Posted by
Lee Cross
What protocol can I use for the pre-fixation permeabilization of...
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.
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.
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
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.
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 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.
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!
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!
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.
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.
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.
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.
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!
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!
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'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!
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.
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.
More
VOTE
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
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
More
VOTE
I am using http://www.biolegend.com/permeabilization-wash-buffer-10x-1497.html followed by http://www.bdbiosciences.com/ptProduct.jsp?prodId=8335&key=fixation+buffer~CATALOG_TYPE:Buffers/Solutions¶m=search&mterms=true&from=dTable
or milder fixator depending on assay
I am using http://www.biolegend.com/permeabilization-wash-buffer-10x-1497.html followed by http://www.bdbiosciences.com/ptProduct.jsp?prodId=8335&key=fixation+buffer~CATALOG_TYPE:Buffers/Solutions¶m=search&mterms=true&from=dTable
or milder fixator depending on assay
More
VOTE
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.
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.
More
VOTE
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.
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.
More
VOTE
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!
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!
More
VOTE
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.
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.
More
VOTE
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.
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.
More
VOTE
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!
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!
More
VOTE
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'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!
More
VOTE