As noted above, fixation will compromise the membrane integrity to some degree or other, with aldehyde fixatives tending to leave the membrane more intact than fixation with agents like acetone, ethanol or methanol. However, one can do some simple labeling at the light microscopic level to rapidly assess just how badly compromised the membrane is in your cells. Careful direct assessment of membrane integrity will require electron microscopic observation.

if you need a quick assessment of how intact a cell membrane is, trypan blue and other exclusion dyes are excellent indicators and often used to quickly assess plasma membrane integrity (most often used with live cells). If the cells exclude the dye, the membrane is intact; if the dye enters the cell, the membrane has been permeabilized. Another quick way to roughly assess membrane integrity is to use fluorescent lipophillic dyes such as DiI or DiO. These dyes intercalate into the membrane and will outline a cell if the plasma membrane is intact. these are available in several different formulations. Another approach to get some rough idea of the integrity of the plasma membrane is by doing labeling with a fluorescently tagged wheat germ agglutinin (WGA), which binds to a carbohydrate moiety found in the glycocalyx of the plasma membrane. You can treat WGA as if its a secondary antibody and incubate your cells with it diluted 1:20-1:100 in PBS ( must contain Ca++!!) for 30 minutes or so at room temp, rinse in fresh PBS a couple of times, then look on the scope.

We, and many others, routinely use 4% paraformaldehyde in 0.1 M HBSS or cacodylate buffer for 30 minutes to fix cell cultures, which gives pretty good preservation of all cellular compartments, including the plasma membrane. To get reliable immunolabeling of intracellular structures, we need to permeabilize the plasma membrane. To do this we utilize 0.1-0.5% Triton X-100 in our blocking buffer (10% normal goat serum + 5% BSA + 0.1% fish gelatin + 0.5% TritonX-100 is our standard for labeling cultured cells, frozen sections and paraffin sections), which we also use to dilute our primary and secondary antibodies. Other detergents also can be used.

If you are trying to label plasma membrane proteins that are especially sensitive to fixation or present in low abundance, like many cell surface receptors, so that even a little loss of antigenicity causes loss of staining, try using a more dilute fixative (1-2% paraformaldehyde) and a short fixation time. We found that fixing longer than 5 minutes caused a major loss of sodium channel labeling in some of our experiments, and a number of neurotransmitter receptors are also very sensitive to length of fixation, too.

If paraformaldehyde fixation simply won't work for you, carbodiimide fixation might might be worth a try (although in our hands the tissue preservation wasn't very good) or a fixative on the market called Prefer that a number of my colleagues have used that shows improved preservation of antigenicity for some, but not all, antigens.

Hope some of this is helpful. Good luck with your studies.

Dave Sherry

There are two possibilities for fixation:

1-organic solvents like methanol and acetone that remove remove lipids and dehydrate the cells. They cause the proteins to localize on the cellular structures.

2-cross-linking reagents like para-formaldehyde. They lead formation of intermolecular bridges through free amino groups. This creates a network of linked antigens. Cell structure is better preserved than organic solvents, but some antibodies do not work on these modified peptides. May cause auto-fluorescence in some tissues.

How do you want to check whether a part of the membrane is gone - to observe this u most likely need high-resolution.

And btw, also PFA will make holes in your membrane (@Korcan), like methanol etc.

And to label any antigen inside your cell you need to permeabilize anyhow your membrane.

Can you maybe specify your question? what do you want to look for?

This is just me speculating, but I would think if you've fixed cells and the membrane is still intact, then you should not be able to stain the cells without permeabilizing first. If you have no issues with staining, than the membrane may be compromised.

Why is it that you want to determine if the membrane is still intact? This will depend on your fixation reagent. As stated above organics will remove membrane lipids, although to different degrees. However, if the protein you wish to study is cytosolic, there is a good chance that the protein will diffuse out of the cell if using organics for fixation. One of the most reliable methods for fixation and permeabilization is 4% paraformaldehyde, and 0.1% Triton X-100 in PBS (30 minutes at RT).

As noted above, fixation will compromise the membrane integrity to some degree or other, with aldehyde fixatives tending to leave the membrane more intact than fixation with agents like acetone, ethanol or methanol. However, one can do some simple labeling at the light microscopic level to rapidly assess just how badly compromised the membrane is in your cells. Careful direct assessment of membrane integrity will require electron microscopic observation.

if you need a quick assessment of how intact a cell membrane is, trypan blue and other exclusion dyes are excellent indicators and often used to quickly assess plasma membrane integrity (most often used with live cells). If the cells exclude the dye, the membrane is intact; if the dye enters the cell, the membrane has been permeabilized. Another quick way to roughly assess membrane integrity is to use fluorescent lipophillic dyes such as DiI or DiO. These dyes intercalate into the membrane and will outline a cell if the plasma membrane is intact. these are available in several different formulations. Another approach to get some rough idea of the integrity of the plasma membrane is by doing labeling with a fluorescently tagged wheat germ agglutinin (WGA), which binds to a carbohydrate moiety found in the glycocalyx of the plasma membrane. You can treat WGA as if its a secondary antibody and incubate your cells with it diluted 1:20-1:100 in PBS ( must contain Ca++!!) for 30 minutes or so at room temp, rinse in fresh PBS a couple of times, then look on the scope.

We, and many others, routinely use 4% paraformaldehyde in 0.1 M HBSS or cacodylate buffer for 30 minutes to fix cell cultures, which gives pretty good preservation of all cellular compartments, including the plasma membrane. To get reliable immunolabeling of intracellular structures, we need to permeabilize the plasma membrane. To do this we utilize 0.1-0.5% Triton X-100 in our blocking buffer (10% normal goat serum + 5% BSA + 0.1% fish gelatin + 0.5% TritonX-100 is our standard for labeling cultured cells, frozen sections and paraffin sections), which we also use to dilute our primary and secondary antibodies. Other detergents also can be used.

If you are trying to label plasma membrane proteins that are especially sensitive to fixation or present in low abundance, like many cell surface receptors, so that even a little loss of antigenicity causes loss of staining, try using a more dilute fixative (1-2% paraformaldehyde) and a short fixation time. We found that fixing longer than 5 minutes caused a major loss of sodium channel labeling in some of our experiments, and a number of neurotransmitter receptors are also very sensitive to length of fixation, too.

If paraformaldehyde fixation simply won't work for you, carbodiimide fixation might might be worth a try (although in our hands the tissue preservation wasn't very good) or a fixative on the market called Prefer that a number of my colleagues have used that shows improved preservation of antigenicity for some, but not all, antigens.

Hope some of this is helpful. Good luck with your studies.

Dave Sherry

Thanks everyone. So, I already understand how all the various fixation methods work and how to permeabilize and all that. The issue is is that people say Methanol removes all the lipids, but that is not necessarily true. I will try to stain the membranes with a lipophilic dye or labeled phospholipids to see if the Methanol (+triton) will remove the PL. I was just having troubles deciding on which dye/label to use to test this, but I think I have it now.

Thanks again.

Dale