How to See Your Proteins of Interest
A short guide to immunofluorescence (IF) experiments – from experimental design to imaging
Visualizing your protein of interest so you can track it and understand it’s position within the cell or subcellular stuctures is an important part of life science research. The problem is, proteins are generally very small and are difficult to see under a microscope. One method for visualizing proteins under the microscope involves using antibodies tagged with fluorescent proteins to bind to your protein of interest. This technique is called, immunofluorescence (IF) microscopy.
We joined forces with Proteintech Ltd. to bring you this resourceful guide on how to stain and image your proteins of interest using immunofluorescence (IF).
7 Steps to great immunofluorescence
IF is based on staining cells with antibodies raised against a target protein that is directly conjugated with a fluorochrome or used together with fluorochrome-conjugated secondary antibodies. IF is a powerful and widely used research technique and the IF results depend greatly on the sample preparation, antibodies, and staining conditions. Here we present a guide to IF experiments, including a brief experiment design, fixation, permeabilization, antibody staining, and the mounting step.
1. Experiment design
It is important to define the research question that the immunofluorescence experiment is intended to answer. When studying the subcellular localization of a target protein, it is vital to use common markers of cellular compartments (Figure 1). Additionally, we need to remember to use quality control markers for our staining procedure. We recommend the routine use of a nuclear counterstain, which can be added to a secondary antibody solution (e.g., DAPI, propidium iodide, Hoechst 3342, or Draq5 – depending on the microscope configuration: light source, wavelengths, and filter sets).
For IF staining we need appropriate controls:
- The unstained sample in order to determine the autofluorescence background signal.
- A sample stained only with secondary antibody to determine the threshold of the background fluorescence signal.
- Tissue/cell type control for those known to express and not express, if possible, the antigen of interest.
- Perform each staining separately; this applies to multiple staining to ensure no cross-reaction and appropriate labeling (Figure 2).
2. Immunostaining cultured cells
Cells are usually seeded on glass coverslips. Strongly adherent cells, e.g. fibroblasts, are able to grow directly on these. Non-adherent and weakly adherent cells (e.g., HEK293 cells) require the glass surfaces to be pre-coated with agents promoting cell attachment such as poly-lysine, collagen, laminin, or gelatin. A similar coating is often used for tissue slices to ensure their attachment to the slide during multiple washing steps. Cells should be at ~50% confluence at the fixation time to yield optimal staining, which allows easy focusing on the specimen during imaging. Too high a cell density may make it difficult to resolve subcellular structures but may also at times be essential, when examining polarized cells that require high density.
3. Sample fixation
In immunofluorescence, cultured cells need to be fixed with antibodies before staining in order to preserve their internal structure. There are two main classes of fixatives: organic solvents and aldehydes.
Organic solvents, such as methanol, ethanol or acetone, dehydrate samples and precipitate proteins, making them accessible for antigen binding. Organic solvents preserve proteins within subcellular structures but also partly wash off lipids and water-soluble components. This means that samples often do not need post-fixation permeabilization to stain intracellular proteins but at the same time organic solvents are not recommended for samples that are intended to be examined by electron microscopy (e.g., correlative light electron microscopy (CLEM)).
Acetone is a stronger dehydrating agent than methanol and is often used in the histological preservation of tissues. Methanol works best for frozen samples and is also often used for fixing cultured cells.
Aldehydes (formaldehyde and glutaraldehyde) fix samples by creating covalent bonds between proteins. Due to their cross-linking properties, they are especially recommended for cytoskeletal components, mitochondrial and nuclear proteins, as well as proteins associated with membranes. The most commonly used protocol involves fixing a sample at room temperature for 10-20 minutes with 2-4% (w/v) paraformaldehyde (Figure 3). Glutaraldehyde is able to cross-link proteins at larger distances than paraformaldehyde but penetrates samples more slowly than paraformaldehyde due to its larger size. Glutaraldehyde is especially recommended for electron microscopy. A common post-fixation step involves quenching the excess of aldehydes in samples to reduce auto-fluorescence (e.g., incubation with glycine). Samples fixed with aldehydes require permeabilization of membranes with detergents prior to staining intracellular proteins.
The choice of fixative pre-determines the permeabilization method and is quite important.
Samples fixed with organic solvents generally do not require permeabilization with detergents and samples can be directly used for antibody staining.
Samples fixed with aldehydes do require permeabilization if the target protein localizes intracellularly. Mild detergents (e.g., saponin, digitonin, or leucoperm) can permeabilize the plasma membrane, allowing efficient staining for cytosolic proteins. Stronger detergents (e.g., NP-40 or Triton X-100) permeabilize the cell membrane as well as intracellular membranes and are therefore suitable for staining nuclear proteins and intramembrane proteins (e.g., ER lumen, Golgi apparatus, or mitochondrial proteins). Alternatively, SDS can be used for permeabilization. SDS partially denatures cross-linked proteins, which often leads to the exposure of inaccessible epitopes and enhances staining for some protein targets.
Using a mild detergent for the permeabilization step (e.g., 0.5-1 mg/ml digitonin prepared in DMSO), try to incubate cells for 10-30 min at room temperature. When working with strong detergent (e.g., 0.1-0.2% (w/v) Triton X-100 prepared in PBS), incubate cells for 10 min at room temperature (Figure 4).
Blocking is an important step that helps to reduce background staining. A blocking agent, such as bovine serum albumin, skimmed milk or serum, is associated with and saturates non-specific binding sites. If serum is used, it is crucial to use one from a species different to that from which the primary antibody was raised, to avoid cross-reactivity. Usually, a one-hour incubation at RT or overnight at 4C with 1-5% blocking agent solution is sufficient.
6. Antibody staining
Using too high an antibody concentration can generate high background signal, while too low a concentration may give no staining at all. The correct antibody concentration depends on a number of factors, including antibody affinity, target protein abundance, and the accessibility of epitopes.
We recommend a series of dilutions, a titration experiment, between 1:20 and 1:1 000 for purified primary antibodies at a stock concentration of 0.2-0.5 mg/ml, which usually covers the optimal range.
It is possible to use antibodies raised against a target protein that is directly conjugated with a fluorochrome (direct labeling) or to perform a two-step staining with an unconjugated primary antibody and fluorochrome-conjugated secondary antibody (indirect labeling). The latter setup is more commonly used and usually offers increased sensitivity due to signal amplification. However, it requires additional washing between antibody incubations to remove excess unbound primary antibody. For co-staining with several antibodies, these need to be raised in different animals to avoid cross-reactivity of secondary antibodies.
Mounting medium prevents sample drying and allows for the long-term storage of a stained specimen. It is important to choose an agent that reduces photobleaching and has a refractive index similar to the objective used for imaging. Sealing the specimen with paraffin wax or fingernail polish adds additional stability and prevents samples from drying.
Dr. Karolina Szczesna
Senior Product Manager and Technical Support Proteintech Ltd.