Cell Structure and Analysis: ICC, IHC, and IF Support—Troubleshooting
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There can be many causes, including insufficient blocking, too high a concentration of the primary or secondary antibody, or degraded primary or secondary antibody. A “no-primary antibody” control can help determine if the secondary antibody is at fault. Otherwise, we recommend trying more stringent blocking or lower concentrations of primary and secondary antibodies.
First, examine unstained/unlabeled tissue under all filter sets to determine that this is not due to endogenous autofluorescence. This is particularly a problem with paraffin sections. If the control still shows this autofluorescence, it may be reduced by washing 3 x 10 min with 1 mg/mL sodium borohydride prior to blocking and labeling.
What may be happening is non-specific binding of the secondary antibody due to dye charge, for example, where the negatively-charged dye is attracted to positively-charged cellular components. To block this, use Image-iT™ FX Signal Enhancer (Cat Nos. I36933 and R37107), which blocks non-specific binding due to charge interactions between the dyes on conjugates and cellular components.
We recommend trying single-color controls, where you attempt to detect one primary antibody with the wrong secondary antibody, and vice versa, to check if there is cross-species labeling. They should be negative. Check the cross-adsorbance species specifications in the product manual of the secondary antibody to insure that the secondary antibodies have been cross-adsorbed against the other species of primary antibody. Another possibility is spectral bleed-through of one dye into the other wavelength. A single-color control (with the correct pairing) can be used to check for this.
Some labels, including some antibodies, are of low-enough affinity that they can come off over time during storage of the labeled slides. To slow this off-rate, samples can be post-fixed with formaldehyde for 5–15 min, after the secondary antibody, to cross-link the secondary antibody in place. The sample should also be mounted in a hardening mounting medium, such as ProLong™ Diamond Antifade Mountant, as the hardening mountant slows diffusion of the secondary antibody. Finally, after the mountant has fully hardened, the slide can be stored cold, preferably at –20°C, to further slow any dissociation.
First, make sure that your cells are mammalian and non-blood derived. If the cells have low transduction efficiency (i.e., only a few cells are labeling or are labeling only dimly), first try increasing the particle number per cell. The next thing to try is to use the BacMam Enhancer Solution. If, however, your cells show labeling, but the localization seems to be too generalized or ubiquitous within the cell, then you should reduce the particle number per cell. The manual gives a guideline and formula for calculating particle number per cell.
Regardless of the type of live-cell indicator dye (e.g., calcium indicators, pH indicator, metal ion indicators), make sure there is no serum during the loading step, which can prematurely cleave dyes with AM esters and bind dyes non-specifically. Always optimize the dye concentration and staining time with a positive control before you run your test samples, to give the best signal-to-background. Always run a positive control with a buffer containing free ions of known concentration and an ionophore to open pores to those ions (for instance, for calcium indicators like Fluo-4 AM, this would include a buffer with added calcium combined with calcimycin, or for pH indicators, buffers of different pHs combined with nigericin). Reactive oxygen indicators, such as CellROX™ Green or H2DCFDA would require a cellular reactive oxygen species (ROS) stimulant as a positive control, such as menadione. Finally, make sure your imaging system has a sensitive detector. Plate readers, for instance, have much lower detector efficiency over background, compared to microscopy or flow cytometry.
Regardless of which dye you use—tetramethylrhodamine, methyl ester (TMRM) or MitoTracker™ Red FM—untreated cells will fluoresce. It’s just that cells with reduced mitochondrial membrane potential will fluoresce less. It is the degree of change which is important. JC-1 dye not only changes intensity, but has a ratiometric spectral change in excitation and emission. It is very important to have an untreated control as well as a positive control treated with a mitochondrial membrane potential destabilizer, such as CCCP or FCCP. These dyes are only for use with live cells, as the signal will not be retained to the same degree with fixation.
When cells and tissues are treated with solvents such as xylene or acetone (for example during deparaffinization of tissue sections), it affects the F-actin in a way that prevents phalloidins from binding. Cryosections, which are not typically washed with organic solvents, can be used instead of paraffin, or anti-actin antibodies may be used.
All fluorescent dyes will fade, or “photobleach,” to at least some extent when exposed to strong light at the wavelengths they absorb. Here are some causes for photobleaching and ways to fix the problem:
Cause of photobleaching
Generation of free radicals and singlet oxygen
Use an antifade reagent, which has antioxidants and free radical scavengers:
-For live-cell imaging of fluorescent dyes and proteins, we recommend ProLong™ Live Antifade Reagent which can be added to the cell media or buffer. ProLong™ Live Antifade Reagent can significantly increase the stability over time for reagents as well as fluorescent proteins, like GFP, without affecting cell health, for up to 24 hours.
-For immediate analysis and short-term storage of fixed samples, we recommend SlowFade™ Diamond Antifade Mountant (which stays liquid and can be used for immediate viewing and then disposal of the sample within a day).
-For long-term analysis of Alexa Fluor™ dyes in fixed samples, we recommend a curing mountant, such as ProLong™ Diamond Antifade Mountant (which slows movement of free radicals).
-For long-term analysis of all dyes and fluorescent proteins in fixed samples, we recommend ProLong™ Diamond Antifade Mountant (which hardens for archiving of slides).
Dye is particularly sensitive to fading
-Choose a more photostable dye, such as many of our Alexa Fluor™ dyes
- Use a counterstain with which you can select and set up your image field, then switch to the dye of interest to image.
- Reduce light exposure, for example by reducing laser power or using neutral density filters.
- Minimize the viewing time of labeled sample, and close shutter when not viewing.
- Use an objective with a lower numerical aperture, such as a lower-power objective.
Here is a good guide to choosing an antifade reagent.
First, make sure you are exciting and detecting the dye in the appropriate wavelengths. Next, try optimizing the dye concentration with your controls, as well as the dye staining time. Our manuals have some guidelines. If you are using a live-cell system, we also recommend washing out any unreacted dye for many of our products to reduce background fluorescence, or adding a background suppressor such as BackDrop™ Suppressor ReadyProbes™ Reagent. Check your instrument settings; some plate readers, for instance, may have a means of adjusting the gain setting to get the best signal-to-background. Finally, some dyes are better than others for degree of change upon ion detection. Contact Tech Support by sending an email to email@example.com if you would like to discuss dye options.
Bubbles may be removed by one of two methods:
- Place the amount of ProLong™ antifade reagent/mounting medium mixture you wish to use on your sample (plus a little excess) in a microcentrifuge tube. Close the cap and centrifuge this aliquot using a tabletop microcentrifuge (speed from 7, 000 to 13,000 rpm). Bubbles should move to the top and these bubbles may be aspirated using a pipettor/pipette tip.
- Unscrew the lid of the bottle/vial containing the ProLong™ antifade reagent/mounting medium mixture to make it loose, but do not remove the lid. Place the entire bottle/vial into a vacuum flask, using a faucet aspirator (faucet T-tube). Apply a vacuum (water running through the faucet) and allow vacuum aspiration to occur from 10 to 20 minutes to degas the mixture.
To avoid the formation of bubbles on a sample or to remove bubbles:
- Before pipetting the desired amount of ProLong™ antifade reagent/mounting medium mixture for mounting, set the pipettor for a slight excess volume. When pipetting up the mixture, do not pipette up the complete amount, but lift up the pipette tip from the bottle with the pipettor not yet up to full volume. This prevents the aspiration of bubbles into the pipette tip.
- Bubbles trapped during application of the coverslip: When placing your coverslip onto your drop of ProLong™ antifade reagent/mounting medium mixture, place the coverslip at a slight angle then, gently lower the coverslip. If the coverslip is lowered flat onto the sample, or lowered too quickly, bubbles can be trapped.
- Bubbles trapped in tissue: One problem with tissue sections, particularly cryosections is that air can get trapped within and under the section. Upon mounting, bubbles are not observed but as the mountant hardens, it compresses the sample slightly, forcing air out of the section. This leads to microscopic bubbles forming over the section, trapped within the mountant. To avoid this, degas the tissue sample prior to mounting. Place the sections submerged in buffer or blocking solution, into a vacuum chamber and expose the sample to the vacuum. This will degas the sections and buffer. Remove the sample from this degassed buffer and mount.
- If ProLong™ antifiade reagent–mounted samples have already cured but have bubbles, you can un-mount your sample by placing the slides into PBS (Coplin jar or a Petri dish filled with PBS). The ProLong™ antifiade reagent will swell and the coverslip will slide off or can be gently removed manually. You can then re-mount with a new aliquot of ProLong™ antifade reagent/mounting medium mixture.
First, make sure you have both a negative (untreated) and positive (ROS-induced) sample to compare. A good positive control can be the use of 100 µM menadione for one hour or 50 µM nefazodone for 24 hours. H2O2 can also be used, though it does not work well for CellROX™ dyes. Some dyes, such as H2DCFDA, require esterase cleavage, so don’t incubate in the presence of serum (which contains esterases that can prematurely cleave the dye). If your positive control does not show significant change compared to the negative control, try increasing the concentration and label time for the dye. Our manuals give starting recommendations. Be sure to image your live cells as soon as possible. Only two dyes (CellROX™ Green and CellROX™ Deep Red) are retained with formaldehyde fixation. Finally, make sure you are using filters and instrument settings to match the excitation and emission spectra of the dye.
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