Immunofluorescence is a powerful technique that uses fluorescently labelled antibodies to detect target antigens, but the choice of fixation method can make a big difference to the results obtained.F1-760-2_rgb

When we fix cells, the goal is to maintain the structure as close to the native state as possible, but to also still allow antibodies to access their antigens.

There are two main groups used for fixation for immunofluorescence. These are the aldehyde fixatives and organic solvents. In many cases the cross linking method using aldehyde fixatives is preferable as it preserves the cell structure better, but there can also be a reduction in antigenicity, meaning that this is not always the case.

This is because the antigens themselves are also cross linked, preventing your antibody from recognising the antigens. To avoid problems due to crosslinking, you can use organic solvents or a permeabilization step after fixing with aldehydes.

In this blog post we are going to look at the main differences you can expect when using two of the most common organic solvents – methanol and acetone.

First, how do the organic solvents work?

Methanol and Acetone act by dehydrogenation and protein precipitation, thereby fixing proteins. This means the cells become instantly permeabilised.

Starting with methanol, we are going to give you some positives and negatives for using the organic solvents as fixatives and details on when you should be using them.


Methanol is most commonly used to fix frozen sections, cell culture cells or smears.

What it’s good for

Preservation of cellular architecture.

Frozen tissue sections and cells.

Stabilising secondary structure of proteins.

Avoiding problems with formaldeyhde crosslinking.

And what it’s bad for

Fluorescent proteins

Tissue morphology.

Strong effect on many epitopes

Water soluble and lipid components can be lost.

Affect the tertiary structure proteins

When should I be using Methanol?

You should be using Methanol when you are trying to visualise antigens on microtubules.

Anything else I should know?

Methanol is often used at -20C. The reasons for this are diverse, ranging from clearer immunofluorescence images to greater control over the process, as fixing using methanol is a quick process and if you didn’t use it at this low temperature then you might end up with not enough lipids to hold any structure.



A strong dehydrating agent, it can cause irreversible precipitation of tissue proteins.

What it’s good for…

Faster procedure

Sometimes less damaging to epitopes.

And what it’s bad for…

Fluorescent proteins

Soluble and lipid components are lost.

When should I be using Acetone?

For better histological preservation than Methanol.

To conserve epitopes to a higher degree.

Anything else I should know?

Acetone will permeabilize and no other permeabilizing step needs to be taken.

Acetone should also used at -20.


How about using Methanol and Acetone together?

Its also worth noting that a combination of methanol and acetone can be used due to acetone being less damaging to epitopes. Acetone is used on snap-frozen tissues, precipitating them, and then methanol is used to fix.

Sometimes Acetone is used after Methanol as a further dehydration step.

What are your thoughts and experiences on using Methanol and Acetone immunofluorescence? Comment below or tweet us at @CiteAb.

– Eleanor and the CiteAb team


If you want to find out more, here is some suggested further reading:

Browse antibodies for use in IF on CiteAb [Link]

R&D Systems: Appropriate fixation of IHC/ICC samples [Link]

Bitesize Bio: Cell and tissue fixation 101 [Link]

Science Lab: How to prepare your specimen for Immunofluorescence [Link]

Cytometry@Leeds: Optimising cell fixation conditions for Immunofluorescence [Link]