Sample preparation for immunofluorescence presents researchers with a range of technical obstacles they must navigate to acquire clear, crisp and informative images. Getting conditions right can mean the difference between overcoming these hurdles and obtaining publication quality data or failing at one or several of these steps and ending up with no signal at all – or worse: acquiring confusing false positives. Here, we briefly discuss sample preparation considerations for better fluorescence microscopy results.
Cell fixation
The goal of fixation is to preserve cell morphology as close to its native state during often rigorous immunostaining protocols. There are two main categories of fixative to choose from, each with a variety of associated pros and cons.
Aldehydes
Aldehyde reagents crosslink free amine groups together, forming intermolecular bridges to create a network of linked antigens. They are an excellent choice for preserving cell morphology, and are especially suited to the visualization of membrane bound proteins. However, their mechanism of action – the covalent alteration of molecular structures – may ultimately reduce target antigenicity. This risk can be mitigated by avoiding prolonged fixation times where possible. Incubation in a 4% formaldehyde solution for 15-20 minutes at room temperature is a popular fixation approach for cell samples. Tissue samples can be trickier, as they generally require longer fixation times – trial and error, in addition to reduction of the tissue section size, may be necessary in such circumstances.
Organic solvents
Organic solvents, such as methanol and acetone, have a dehydrating effect on the cell, which precipitates proteins, fixing them in their cellular context. They are generally excellent at preserving cellular architecture and negate the requirement of any subsequent permeabilization steps – you can go directly to your staining protocol following sample fixation. Application is also straightforward (no involved preparation techniques, unlike aldehydes); ice-cold methanol (-20C) can simply be applied to cells for 10-20 minutes, before being rinsed away with buffer. The (slightly more complex) combination of 1:1 chilled methanol and acetone may yield even better results – acetone improves sample permeabilization without being as harsh on epitopes as methanol.
Despite the pluses, there are a few important contraindications of organic solvent fixation to be aware of. They remove a lot of small, soluble molecules and lipids in the fixation process, making them unsuitable for use in certain scenarios. They also denature genetically encoded fluorescent proteins so are a definite ‘no-no’ if you are counterstaining with these fluorescent probes.
Permeabilization
Antibody proteins are too large and ionic in nature to pass through cell membranes without prior permeabilization – an essential measure if aldehyde fixation has been employed. Several detergents are available that offer differing degrees of membrane disruption and levels of access to subcellular compartments.
Triton-X-100 is a non-ionic detergent widely used for permeabilization as it efficiently dissolves cell membranes without disrupting protein-protein interactions; it also grants access to the nuclear interior. It is typically used at 0.1 %-0.2 % in PBS buffer for 10 minutes. Longer incubations or higher amounts of the detergent are not recommended due to its somewhat harsh nature. Alternatively, digitonin or related saponin compounds which are milder detergents can be used in place of Triton-X-100 to disrupt membranes.
Buffer selection
For the most part, phosphate buffered saline (PBS) is the buffer of choice for most fluorescence imaging experiments; it is isotonic, does not disrupt cellular structure and maintains pH near physiological levels. However, if you’ve obtained weak signal at the end of your experiment it could be that your antibody is not happy in PBS. Antibodies are optimized to work in the sodium rich environment of the blood, yet the intracellular environment predominantly contains potassium. Trying out alternative buffers post-permeabilization could improve antibody signal.
Blocking
Blocking prevents primary and secondary antibodies from binding non-specifically to your sample, by masking potential sites of weak, generic interaction. Bovine serum albumin (BSA) is commonly used for this purpose, but fetal calf serum (FCS), casein (or a solution of non-fat dry milk), gelatin, or normal serum (which must originate from same species as your secondary antibody) can also be used.
If your sample is permeabilized, it is also advisable to include a small amount of detergent to out-compete antibodies for hydrophobic interactions.
Antibody options
Antibodies can make or break your immunostaining protocol. It pays to know the benefits and limitations of options like direct and indirect staining, the use of monoclonal vs polyclonal antibodies, and which secondary antibodies to use.
Direct vs indirect staining
Direct staining, in which the primary antibody carries its own fluorophore, enables you to bypass several steps in the fluorescence staining protocol. Indirect staining is often used when detecting low abundance proteins as it enables amplification of signal. Indirect staining is more adaptable and can be cost effective in certain situations.
Primary antibodies
Monoclonal antibodies originate from individual B cell clones and only bind strictly defined epitopes. Polyclonals are a heterogeneous mixture of antibodies from different B cell clones with different specificities and affinities for the same target protein. You can exploit their inherent differences to the advantage of your experiment. Precision work, like staining c-termini, may be better with a monoclonal; whereas polyclonals, especially those raised against larger protein segments, would be better suited to targets with low copy numbers. Regardless of whether you are using monoclonal or polyclonal antibodies for staining, it is advisable to carry out antibody titration with each new antibody batch based on the vendor recommendations. Most commercially available antibodies have sufficient titer to bind targets in 30-60 minutes at room temperature but some targets may require an overnight incubation at 4°C.
Secondary antibodies
It is important that secondary antibodies not only recognize a primary antibody’s host species, but also its isotype. Pre-absorbed secondary antibodies reduce the risk of cross reactivity with non-specific targets. Pre-adsorption is an additional processing step where secondary antibodies are passed through a column matrix containing immobilized serum proteins from potentially cross reactive species. They are highly recommended for multi-color experiments when several primary antibodies and their corresponding secondary antibodies are used simultaneously. Pay attention to avoid overlapping excitatory wavelengths that could result in suboptimal data.
Washing
Excess reagents and antibodies need to be removed following each incubation to prevent any interference with the next step of your protocol. Two to three cycles of buffer aspiration and additions performed in sequence should be sufficient, with the final step being an aspiration to remove any fluid prior to the next reagent. It is vital at this stage to move in a timely manner; leaving your sample exposed for too long without buffer will cause it to dry out, thus ruining your experiment.
Counterstaining
Obtaining strong, specific signal from a protein of interest is a welcome outcome of any immunofluorescence imaging experiment; however, specific signal without context, no matter how clean and clear, can be difficult to interpret or even meaningless. Unless there are good reasons for not doing so, you should always include a counterstain in your experiment. Nuclear counterstains, like DAPI and Hoechst 33342, are popular options. Other cellular structures you may consider counterstaining are the actin cytoskeleton using fluorescently-labeled phalloidin, or the plasma membrane using fluorescently-tagged wheat germ agglutinin (WGA). Unlike antibodies, counterstains usually do not cross react with one another and can often be incubated together in a single step (the only limitation being buffer compatibility).
Sample preservation
Taking measures to preserve your imaging experiment is just as important as each of the preparation steps above in terms of obtaining quality data. Mounting medium helps preserve the sample and raises its refractive index (which improves performance with oil objectives); it also often contains agents that scavenge free radicals and reduce photobleaching. After sealing coverslips, store your slides in an opaque, light-tight container at 4°C in case you need to reference them at a later date.
Final words
There is no one-fits-all sample preparation protocol to suit every immunofluorescence scenario. The optimum staining protocol for a nuclear protein will hardly resemble that of one for an intracellular plasma membrane-bound target. There are numerous methods to fix, permeabilize, and stain cells, each with their strengths and weaknesses. Rather than a single strategy you should always rely on, you should tailor each step of your immunostaining procedure to the target or targets you wish to visualize. Always test a number of different techniques in order to optimize results.
Image: Courtsey of BioTek Instruments. Shown is an example of an Alzheimer’s Disease Cellular Model: Red:immunofluorescence targeted to aggregated β-amyloid peptide binding to neuroblastoma cell line; counterstained with Blue: Hoechst stained nuclei; Green: phalloidin stained F-actin subunits (cytoskeleton)
Related Products from: BioTek Instruments