Optimising cell fixation conditions for immunofluorescence

Fluorescence microscopy commonly utilises a technique called immunofluorescence (IF) where antibodies against a primary target are detected using a fluorescently conjugated secondary antibody. This often has the effect of amplyfying the signal from the primary and allows for bright subcellular structures to be observed. However, not all antibodies work as efficiently as they should! I'll discuss fixation techniques here as a way of maybe approaching your IF experiments if things aren't working so well.

People often use paraformaldehyde (PFA) as a fixative for IF. PFA has the effect of cross-linking aldehyde groups and effectively fixes your cells as a jelly. PFA generally does not effect the conformation of proteins and often monoclonal Abs prefer PFA fixation. This jelly then needs to be permeabilised to enable antibodies to penetrate and find intracellular antigen targets. We usually follow PFA fixation with detergent such as TritonX100 or saponin. PFA concentrations of 3-4% are often used.

However, PFA can cause some permeabilisation depending on the cell type (~20% with HeLa cells fixed for 20 mins in 3% PFA). One potential way around this if you only want to label the outside of cells is to reduce the PFA concentration to 1 or 0.5% and fix for 5 mins. FACS analysis can prove to be an effective method to titrate PFA fixation condition.

In the past I've used a mix of 3% PFA with 0.2% NP40 detergent as a fixative! Fixing and permeabilsing at the same time! I recall it gave really nice clean samples with very little background.

Methanol is also often used to fix cells, especially if microtubules are the desired target. Fixing cells in -20degC methanol rapidly fixes cellular structures and allows the cytoskeleton to be effectively preserved. However, my experience is that room temp methanol is effective too! Methanol acts to dehydrate cells and removes the membranes and lipids. Methanol 'denatures' proteins upon fixation by effectively precipitating them out. When you fix with methanol there is no need for a permeabilisation step.

Acetone is often used following methanol fixation as an additional dehydration step. In my experience its often not required but it will suit some peoples application. 10-15 mins in methanol followed by 5 mins with acetone is often recommended.

I've used methanol previously as a permeabilisation method following PFA fixation when detecting lamin in the nuclear envelope. I've even mixed PFA at 3% with 50% methanol as a fixative, seemed pretty effective at opening up nuclear antigens - anything is worth trying!

Other fixative commonly used are formalin (also referred to as Sigmafix if brought from, well, Sigma), this is the same as PFA except it contains a little methanol that prevents the formalin turning to formic acid. The presence of methanol will effect your experiments if you want to label the cell membrane alone.

Ethanol is sometimes used as a fixative and effectively can be viewed the same as methanol. Ice cold ethanol is wonderful for accessing nuclear DNA for cell cycle analysis with dyes like DAPI and propidium iodide.

You can even microwave your samples following PFA fixation. This has the effect of permeabilising your cells and can be quite effective at retrieving your antigen. Search for 'antigen retrieval microwave' in Google for some protocols.

So in conclusion, all antibodies have optimal fixation conditions and it's worth exploring them all to get good quality images of your samples. Sometimes if you are labelling with two or three different antibodies they may have different preferred fixation conditions! This is when you have to experiment a little.