Specific staining with high signal and low background is not easy to achieve with immunofluoresence or other fluorescent staining. Below are some common problems and how we recommend solving them. If you have any additions, suggestions, or corrections, please let us know.Troubleshooting immunofluoresence
The following table outlines some common problems, their probable cause, and possible solutions. As with all imaging issues, we recommend discussing any problems you have with expert members of our staff. Identifying these problems may be more subtle than the language below suggests. Please consult us. We're here to help you.
|Cells look flat or like impact craters||cells allowed to air dry||Do not allow cells to dry at any point.
Put coverslips/chambers etc. inside plastic box lined with rolled up wet paper towel during incubations.
|Cells look flat or like impact craters||methanol fixation||Fixation should preserve morphology. Methanol causes membrane to collapse. Find alternative fix such as formalin or glutaraldehyde.|
|Tissue or cells autofluoresce||1. fixation with glutaraldehyde
2. material is autofluorescent
|Quench after fixation and before staining. Try:
1. Incubate with NaIO4 (21.4 mg/ml in PBS) 15 min.; PBS wash.
2. NaBH4 (10mg/ml) in PBS approx. 4 repeated washes 15 min. each; NaBH4 should be dry until immediately before use; PBS wash. [More.]
3. Incubate with glycine; PBS wash.
Screen for autofluorescence before staining.
|Cells look dim; mounting medium looks bright||1. background too bright due to insufficient washing
2. binding too weak, especially with phalloidin or phallocidin
|1. Wash better
2. Check that secondary antibody or other probe is binding tightly enough
3. Better fixation.
|Bright spots of fluorescence appear all over section, cells, or slide||1. Secondary antibody too concentrated
2. Secondary antibody precipitating
3. Nonspecific binding of primary or secondary antibody
|1. Reduce concentration of secondary
2. Filter or spin secondary antibody/fluorophore
3. Do controls for nonspecific binding; if coverslip/slide is speckled, chances are the problem is 1. or 2., not this.
|No labeling on sample even though good on Western blots||1. fixation destroys antigenicity
2. insufficient permeabilization
2. permeabilization washes out antigen
3. the antibodies are dead
4. antibody raised against peptide not whole protein
5. what you think is there, really isn't there
|Cells constantly drifting in and out of focus.||1. Coverslip is not fixed in place, not sealed with nail polish or
2. Cells not adherent to coverslip
3. microscope objective crashed into coverslip
|1. Seal coverslip
2. Use polylysine or other adherence
3. Bring focus down; new oil on objective
|More on quenching aldehyde groups left after
From: "James F. Sanzo" sanzo@AVIGENICS.COM
There are several other methods you can try that quench the remaining free aldehyde groups left after glutaraldehyde fixation (below). However, if you are not necessarily interested in ultrastructure, why not use paraformaldehyde? Aldehyde induced fluorescence is usually much less of a problem with the PFA. You may also want to think about whether the fluorescence you are seeing is indeed due to aldehydes - or if it could be due to some other cellular constituent like lysolipids. As an alternative, you may also want to consider using the longest wavelength your imaging and fluorophore systems will allow. Try looking at the background fluorescence above around 590nm. It may be all you need to do.
For blocking free aldehydes remaining after fixation:
1. 1% (or 150mM) glycine with 0.1% Tween in pbs. Wash well. (glycine binds to free aldehydes)
2. 50 mM NH4Cl in PBS. Incubate for 15 min at RT. Wash well. (see Bendyan)
3. 0.1% Na borohydride in PBS. Apply while fizzing and incubate 3 x 10 min on ice. Wash well. (reduces carbonyls)
4. 0.15 M ethanolamine, pH 7.5. Incubate 30 min on ice. Wash well.
From: Scott Snyder SSnyder@NIAID.NIH.GOV
Probably the best way I have found of getting rid of aldehyde based fluorescence is to get rid of the aldehydes. I often do this by using the bifunctional crosslinking reagent dithio bis(succinimidyl propionate), or DSP for short. This gives comparable structure retention in my hands and essentially no added fluorescence. A good reference for this is Safieko-Mroczka and Bell, Journal of Histochemistry and Cytochemistry v44 No. 6 641-656 (1996).
originally by cbm; last revised 3 June 2002 by mc