Oct 15, 20236 min read

Doing more with less: Fixatives

Updated: Feb 16

Which fixative should I use?

PFA? A kit? Maybe formalin and dish soap?

Answer: It depends!

Honestly, it depends a lot on what you're trying to accomplish. Different fixatives do very different things to your cells. You'll want to consider what matters most, whether it be accessing the nucleus, preserving cell surface epitopes, keeping the scatter profiles intact or even maintaining phosphorylation states. I'll give some examples below of how some commonly used fixatives affect staining for some markers with mouse cells (sorry, no human data today), and provide a rundown of the features of these fixatives.

Before I get to that, a safety note: fixatives are by their nature toxic! The reagents I'm talking about today all contain some version of formaldehyde, which is neurotoxic and carcinogenic. Please use appropriate precautions while working with these reagents, and dispose of them correctly, not down the sink.

Why do we want to fix the cells? Fixatives form covalent chemical bonds between nearby molecules in the cells. This creates a rigid structure that is more stable, preserving the cells for storage or allowing us to do things like punching holes in the membranes without completely disintegrating the cells. So, we can keep the samples for longer before acquisition and we try to stain for molecules inside the cells.

In order to stain intracellular components, we also need a permeabilising agent. These are almost always detergents that serve to wash away lipids from the membranes, creating holes that the antibodies can pass through. In many cases, the "fixative" in a kit will be a mixture of a fixative (e.g., PFA) and a detergent (e.g., triton). For example, BD's Cytofix/Cytoperm, Thermo's eBioscience Foxp3 Fix/Perm buffer, and BioLegend's True-Nuclear buffer. Alternatively, the fixative and permeabilisation agents may be applied sequentially, such as using PFA to fix and then saponin to permeabilise.

In general, mixing the fixative and permeabilising agents into one solution allows for better access to the nucleus and other intracellular compartments, and generally preserves epitopes better for staining (the detergent probably prevents some of the crosslinking). The downside is that you get extensive loss of free-floating cytoplasmic components. This is particularly notable with fluorescent proteins like cytosolic GFP (not membrane-bound GFP or nuclear GFP) and cytokines.

Fixing and permeabilising the cells means the antibodies can now get inside the cell, where there's way more stuff than on the surface. This can be detrimental for staining if the antibody has off-target binding to proteins inside the cell. In some cases, however, the target protein that we think of as a cell surface protein is actually largely inside the cell. CD3 epsilon stains way better if you permeabilise the cell, perhaps due to internalisation or just being transmembrane. CCR7 and other chemokine receptors can be internalise upon binding their ligands, so CCR7 stains much better on cells from lymphoid organs with permeabilisation. Cells that are undergoing stimulation, for example in cytokine staining assays, will exhibit down regulation of many proteins, so even CD4 and CD8 stain much better internally in this setting. It's worth remembering, too, that all the proteins are made first inside the cell, so there will be stores of many surface proteins on the inside.

Here's an example of how intracellular staining for a cell surface protein, CD11b, can improve detection:

The splenocytes here have either been stained without fixation or stained after fixing and permeabilising using the eBioscience Foxp3 Fix/Perm kit. There's an order of magnitude of increase in the signal, plus a slight decrease in the noise. Also, we can now see differential expression of CD11b in different cell types (highest to lowest: neutrophils, monocytes, myeloid DCs, NKs).

Here's how that works out with five different fixatives:

eBioscience Foxp3 Fix/Perm
IC Fixation Buffer (PFA)
Formalin at 0.2% in PBS
Formalin at 2% in PBS
BioLegend True-Nuclear (TN)

PFA doesn't help us at all over unfixed cells. This is because PFA causes noticeable degradation of most epitopes when used at a 4% concentration, and, in my hands, behaves inconsistently at lower concentrations. PFA also isn't very stable over time.

Formalin (formaldehyde) displays time- and concentration-dependent reductions in staining for most of the same epitopes that are affected by PFA (they are nearly the same). However, this can be consistently controlled. For a strong fixation that preserves scatter, fluorescent proteins and cytokines, 2-4% for 30-45min works nicely. To preserve epitope detection while keeping cells intact for intracellular staining, 0.2% for 20-30min is good.

The eBioscience Foxp3 Fix/Perm kit provides a good all-around solution for immune phenotyping as it preserves most epitopes and permits good intranuclear staining. It is not suitable for detection of cytoplasmic fluorescent proteins or most cytokines. Also, the high saponin content of the permeabilisation buffer can slightly increase the background of the cells.

The True-Nuclear kit gives good nuclear staining, and is surprisingly good for detecting Cas9. The permeabilisation buffer gives less background than the eBioscience one. However, this kit isn't compatible with all fluorophores and has some odd effects on some epitopes. Here's a comparison of the eBioscience Foxp3 kit versus the BioLegend True Nuclear kit for some nuclear antigens. I'm using overnight staining for Foxp3, Helios, Ki67, Bcl-6 and GATA-3, while the CD4, CD3, CD19 and viability markers were stained for 1hr prior to fixation. As we can see, the background of all cells has shifted up in the eFluor450 channel with the eBioscience kit relative to the BioLegend one. This occurs from about 430 to 600nm, mainly in the violet. You can reduce this background by using the True Nuclear permeabilisation buffer in place of the eBioscience one, or by using autofluorescence subtraction.

Here are some examples of how these different fixatives affect staining for a few choice markers on mouse splenocytes. In all cases, I've stained for CD4, viability and Foxp3 (using a Thy1.1 surface reporter) before fixation. The plots below are gated on viable CD4 T cells, and I've selected the optimal concentration, which is not the same in all cases. The "Unfixed" cells are also unpermeabilised.

I recommend staining CD69 prior to fixation. Helios is quite sensitive to crosslinking (and without permeabilisation, it can't be detected).

Note that by staining for IgM after permeabilisation, we can detect the rare IgM plasma cells as a CD19-dim IgM-bright population because they are antibody factories.

I mentioned that formaldehyde (formalin) is time and concentration-dependent. Here's how that works using Foxp3 and Helios staining as an example:

Cytokine staining is highly dependent on the fixative used. Using a fixative with an added permeabilising agent usually causes loss of certain cytokines, particularly IL-2, IL-4 and IL-10, due to opening up of the plasma membrane. To retain cytokines inside the cell, the best approach is to fix completely first, then permeabilise. I'll go over best approaches for intracellular cytokine staining in a separate post.

Figure details: Viable CD4+CD3+ T cells after stimulation for 4hrs at 37C with 500ng/ml PdBU, 750ng/ml ionomycin and in the presence of 1ug/ml brefeldin A to block secretion. All staining performed overnight after fixation.

For details on how to best preserve your cytosolic fluorescent reporters (e.g., GFP), I suggest reading this paper:

https://onlinelibrary.wiley.com/doi/10.1002/cyto.a.22451

Summary:

eBioscience Fix/Perm kit: