Cell Imaging Support—Troubleshooting

Here is a page you can go to. Look under the “Cell Imaging Systems” section and follow the link for your EVOS™ imaging system. There you will find the free download link as well as instructions to follow. We recommend you check for updates at least every six months, or if your system seems to have a software glitch of any sort.

If the lens is running up into a sample, this may be an issue with either focusing too quickly and missing the focal plane (if focusing manually) or a problem with the objective calibration (if using autofocus). It is a good idea to calibrate your objectives using the FL Auto calibration slide that comes with the system. Check to see if your objective is a long-working distance (LWD) or coverslip-corrected objective (CC). If coverslip-corrected, it is only for use with very short working distances for imaging through thin coverslips, but not through the slide or through plastics in microplates or culture dishes). If working with high magnification and oil immersion, by eye, move the objective upwards to touch the bottom of the sample and then only move slowly away from the sample for further focusing. 

An objective lens can be seriously damaged by scraping against samples. If this happens, check the objective lens for damage.

Objectives can hit the vessel holder when they are focused too high in the Z axis (up and down). This is a particularly a problem with the EVOS™ FL Auto Imaging System during instrument start-up, when the stage moves during system initiation, or when changing objectives. Coverslip-corrected objectives tend to be wider and flatter at the top of the barrel, which means that they are more likely to run into the edges of the vessel holder, particularly if you are imaging at the edges of the sample container. In those cases, use of that objective for those areas of the container may not be possible. If the objective if “jammed” by the vessel holder, then carefully unscrew the thumbscrews of the vessel holder and lift it straight off the stage, then move the objective downward in focus and toward the center of the stage. It is a good idea to have a shut-down procedure in your lab that includes moving the objectives to the lowest magnification and focusing downward with course focus prior to turning off the instrument for the day.

An objective can be damaged by scraping against the vessel holder. If this happens, take out the objective and examine it carefully for damage, particularly on the lens. 

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.

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.

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. Phalloidin may be used with cryosections, which are not typically washed with organic solvents, 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:

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 techsupport@thermofisher.com if you would like to discuss dye options.

Bubbles may be removed by one of two methods:

1. 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. 
2. 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:

1. 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. 
2. 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.
3. 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. 
4. 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.