Fixation and flow cytometry
Fixation is routinely used in histology and cytology Labs the world over as a way of keeping cells in stasis at a particular point to ensure that, by the time they are examined, they have not deteriorated. This is also something that we often want to do in flow cytometry experiments. It seems like a simple procedure but there are many ways of doing this and, as usual, there isn’t not one universal way to success.
Why should I fix my cells?
There are many reasons why fixing your cells is a good idea:
- It will allow you to run samples when you are ready to and when a cytometer is available. This is of particular benefit if you are doing a time-course assay (or are paying for access to a cytometer!).
- It will be safer because your cells are no longer alive; in fact in some labs fixed samples may be mandatory
- It is convenient and easy
- If the antigen or target you want to look at is intracellular you’ll most likely need to fix your cells in order to permeabilise them allowing the antibody/dye to reach its target.
So, if you want, or need, to fix, what are your choices? Simply put, there are two commonly used classes of fixative – alcohols or aldehydes.
Alcohols, usually ethanol or methanol, are precipitating or denaturing fixatives that coagulate proteins. They will also dissolve lipids meaning that they will create large holes in the cellular membranes (plasma membrane and nuclear membrane). Alcohols are generally good when you are interested in DNA analysis – coagulation of the proteins around DNA allows even access to the dye, leading to a good coefficient of variation (CV) (low data spread). But, as you may have thought, coagulation of proteins can be problematic if your antibody depends on a particular conformation of its target protein to recognise its epitope. So in very many cases alcohol fixation will mean you will be unable to detect your protein even though it is there in the cell! The exception to this will be small epitopes such as phosphorylation-specific antibodies.
Aldehydes are cross-linking fixatives and create bonds chiefly between lysine residues. In this way they keep the structure of most proteins intact, which makes them an ideal candidate for antibody staining as the epitope is more likely to remain intact.
The usual aldehyde fixative used in cytometry is formaldeyde, which will polymerise to become paraformaldehyde (PFA) unless there is a small amount of methanol added to the solution. A common mistake is to say that paraformaldehyde is used to fix cells. In reality, paraformaldehyde is an insoluble white powder that won’t fix anything – the solution we use is methanol-free formaldehyde. For fixation purposes, it should be diluted with PBS to a concentration anywhere between 0.5% and 4%. For a more in depth explanation of the nomenclature of formaldehyde and paraformaldhye see this message on the Purdue University Cytometry website.
Glutaraldehyde is extensively used in electron microscopy, but is not as widespread in flow cytometry. This is because it is more effective at cross-linking that formaldehyde and therefore it is harder for antibodies and dyes to stain structures effectively; it also increases cellular autofluorescence markedly so unless you have a particular reason for using glutaraldehyde I’d stick with formaldehyde or alcohol.
How long will my cells last once fixed?
You’ll be pleased to hear that cells fixed in ethanol are stable at 4oC or –20oC for months. For formaldehyde fixed cells this is reduced to around 1–2 weeks, but, unlike ethanol fixed cells which can happily sit in ethanol all that time, you should spin them out of the formaldehyde after 2-3 hours fixation since prolonged storage in fixative will increase autofluorescence. After this they can be safely stored in PBS,. Increased autofluoresence is something to be wary of with all fixatives and it can be particularly problematic if the target of interest is expressed only weakly. Fixation may also change the light scattering properties of the cells, which may be important to you.
Combining fixatives: the best of both worlds
It is also possible to combine fixatives and a common way of doing this is to first fix in formaldehyde and then post-fix in alcohol; the first fixative locks everything in place in the cells and the second fixatives makes larger holes in the cell allowing easier access of antibodies and probes. It is also possible to use a permeabilising agent such as Triton X-100 or Tween-20 as the secondary step.
A few tips for fantastic fixation
- Order of staining: The order in which you stain and fix may vary depending on the biological question being asked. Sometimes if surface and intracellular epitopes are to be examined, it is best to stain surface antigens first then fix. But beware, some fluorochromes are sensitive to fixation. Fluorochromes such as phycoerythrin (PE) or allophycocyanin (APC) are large protein molecules and will be affected the same way as other proteins by the fixation so try to avoid alcohols. Small fluorochromes such as AlexaFluor488 or FITC are generally unaffected by whichever fixative is used.
- Use cold solutions: There are also a couple of practical tips when fixing that are useful to bear in mind:
- Fixation should generally be done using a cold solution as the process of fixation is exothermic.
- Gently vortex: Cells should be vortexed gently while adding fixative to ensure good penetration of the fixative and reduce cell clumping. And add at least 1ml of fixative solution to a cell pellet.
As long as you are aware of the chemical effects of fixation and of the types of fixative used, you can vary the process according to your needs which can lead the way to a successful flow experiment!
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All the articles are very informative and good content. Thank you.
My untreated cells are too dead by the time I do the flow cytometry. I’m using fitc-annexin v and pi to stain the cells. Should I fix the cells after staining to preserve the cell state?