Immunocytochemistry protocol – version 2.2, 23 Sept 2010

Purpose: To get high-magnification images of cellular and sub-cellular structures through fluorescent antibody staining

Before you begin: It is easiest to get high-magnification/high-resolution images  if you grow your cells on a coverslip, although a chamber slide is OK.  The closer your cells are to the microscope lens, the easier it will be to see them without risking crushing the slide with the lens.  Only use coverslips of thickness 1 – thicker slips (1.5 or 2) make focusing very difficult at high magnifications.

In general HEK293 cells stick poorly to glass – you may get better results with coated glass (poly-L-lysine, collagen or a commercial coating such as is on chamber slides).  HeLa, U2OS and other strongly adherent cells are fine on plain glass.  Plate and transfect your cells so that they are subconfluent prior to fixation – you will see the cell architecture better and have less piling up of cells, also most cells adhere better when not in a single continuous sheet.

If using coverslips, it is easiest to keep them in the dish for the first 3 steps and only take them out to stain with antibody.

Procedure:

1. Fix cells in 4% formaldehyde solution in PBS for 15 mins.  Wash once in PBS.  (Some antigens may be better fixed in methanol, this varies between antigens, methanol fixed cells do not need permeabilisation but stain poorly with phalloidin and other confirmation-specific stains). For mitochondria and other organelles you may find adding 5% sucrose to your fixation buffer better preserves morphology.

2. Permeabilise in PBS, 1% BSA, 0.1% Triton X-100 for 2-5 minutes (Caco-2 and similar cells may take longer, HEKs may come off the glass if left too long); you can also use 0.5% saponin in 1% BSA PBS for 20 minutes if you need to preserve membrane structure, but you will need to do all your subsequent antibody incubations and washes in 0.1% saponin (saponin forms pores in the membrane, but these will wash out after a while).  Delicate membrane structures and compartments are better preserved with saponin, but it will not permeabilise the nuclear membrane.  If you plan on using serum to block your cells, do not use BSA here (it is not required and may increase your background).

3. Wash once in PBS and remove coverslips from plate to parafilm or other support.  If you need to use very small amounts of antibody you can invert coverslips onto a small drop of liquid (e.g. 20-40 μl), otherwise an 18mm coverslip can be easily covered with 100 μl and you are less likely to drop or break coverslips this way.

4. If you are doing multiple stainings or are staining tissue sections or cells with Fc receptors you should block with appropriate serum here.  An appropriate serum is one which does not react with your secondary antibodies, usually the same species as your secondaries.  Block in 2-5% serum for 10 minutes, do not wash after, just suck the liquid away and add antibody.

5. Stain with primary antibody at suitable concentration (between 1:100 and 1:1000) for 60 minutes at room temp.  Dilute antibody in PBS (some people add 0.1-1% BSA to this).

6. Wash well 3 x 5 minutes in PBS.

7. Add secondary antibody (between 1:200 and 1:500). Incubate in the dark for 60 minutes.

8. Wash well 3 x 5 minutes in PBS.  If you are using phalloidin wash just once, add phalloidin for 20-40 minutes, and then wash well 3 x 5 minutes.

9. One-by-one, gently dry coverslips by picking them up, touching the edge to some tissue paper and mount on a slide using aqueous mountant.  Seal edges with nail varnish.

10. Leave in dark to set for 15 minutes before imaging.

If staining with multiple antibodies you can do two things – follow the steps above and repeat steps 5-8 for each primary and secondary antibody pair (recommended, see below); or stain with both primaries, wash, then stain with both secondaries simultaneously.  To do simultaneous stains you must make sure your antibodies do not cross-react and include single-stained controls.

 

Tips and tricks:

A good article on imaging, especially for multiple colour antibody staining is - The good, the bad and the ugly Helen Pearson, Nature doi:10.1038/447138a

Gentle handling is a must – add solutions gently from the side of the slip or slide.

Do not let your cells dry out at any point during staining, this can introduce artifacts and cell shrinkage.

If you have strong signal properly mounted fluorescent staining will keep for 7 days or longer in a dark, cold (4°C) place.  Some expensive mountants (such as Molecular Probes Prolong Gold) allow you to store them much longer than this (I have used slides over a year after they were mounted with Prolong Gold).

Always include single staining controls – you must do this when using two antibodies together for the first time or under new experimental conditions e.g. serum starvation, infection of cells.  Without these controls you will not be able to tell if your antigens are in the same locations, or you have cross-reactions (proteins move around in response to conditions in the cell, without controls you will not known whether you have colocalisation or an artifact). 

If you do not see the same locations in your single-stains as in the double-stained samples you have a problem.  You can reduce the possibilities of generating artifacts with the tips below, but must still have single staining controls.

In general your chances of artifacts are reduced by doing sequential staining (primary A, secondary A, primary B, secondary B) rather than mixing primaries and secondaries.

For multiple staining you need to be sure your secondary antibodies are cross-adsorbed against (at least) your other secondary and primary species.  You can get such high-quality antibodies in any colour from a number of vendors including Invitrogen/Molecular Probes (labeled “highly-cross adsorbed”) and Jackson ImmunoResearch (their anti-IgG donkey antibodies are excellent).  Ideally your secondaries will be from the same species.

Excessive background can be reduced by: blocking with serum from your secondary antibody species; reducing antibody concentrations and/or incubation times; quenching residual aldehyde groups following formaldehyde fixation with 0.1M glycine (pH 7.5) or 0.1% sodium borohydride in PBS (this has to be made fresh).