Long-term Live Cell Imaging tips/tricks

Hello – My project requires me to image cultured cells (macrophages) for up to 72 hours on an epi-fluorescence microscope. Are there any specific tips and tricks that people have that I should consider when doing these experiments? I’m using an Okolab incubator with continuous 5% CO2 supply. Two of the specific issues I have been having is:

  • Medium stability

  • Evaporation
    Any help is appreciated!

  • Maikel


Hi @Maikel! To decrease evaporation, we recommend a layer of mineral oil on top of the culture media. In our humidified Okolab incubators, that’s usually sufficient to counteract evaporation for multi-day experiments at 37*. Make sure the mineral oil completely covers the surface of the culture media.

If the mineral oil isn’t cutting it, or there’s some reason that it’s not feasible with your setup, you’ll probably need to find a way to increase the local humidity at the sample. Does your incubator have a built-in humidity system?


Thank you @Anna! That sounds like a good solution. We do have the humidity system and that definitely does help with a large part of the evaporation, but the sample requires a ‘re-fill’ of medium over time (which is sub-optimal). Is there a specific mineral oil you use?

Mineral oil works very well for reducing evaporation. It can also help to fill any empty wells in a multi-well plate with water.

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We’ve used a few varieties over the years, including some from the drugstore, and they all seem to work fine. The one we have right now is a light mineral oil from Sigma.


Thank you both! That is very helpful. I’ll give it a shot.

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I’ve run man MANY longterm timelapse experiments. As suggested, mineral oil is a great tip, just be sure to layer enough on top so that the entire surface area is covered, in practice I find that warming the mineral oil a bit before hand also makes it easier when layering on top - avoid bubbles as they will mess-up your brightfield/phase/DIC. I used mineral oil from Sigma that is for mouse embryo work. Regarding medium “stability”, some people have luck adjusting buffering capacity by tweaking sodium bicarbonate or supplementation with HEPES (25mM final, pH 7.4), but with a good incubator this shouldn’t be much of a point. For some cell lines I’ve used CO2 independent medium and turned off the CO2, not all cell lines like the CO2 independent medium, it can be mixed with normal medium to ‘wean’ your cells onto it. I assume you’re system uses a modern autofocusing system that is independent of visible fluorescence, if not, be sure not to focus using fluorescent light. What do you mean by “medium stability”?


Thank you for your response, James. I will look into the HEPES and Sodium Bicarbonate. You’re absolutely right our Nikon system has a PFS module that uses infrared. With medium stability I mean that the medium discolors (change in pH) pretty rapidly under the microscope. Now that I’m reading up on it, HEPES might be a great solution for that issue.


For media stability, you can also try media that are not bicarbonate based. For some of our long term imaging projects we use Leibovitz (L-15) media. Check media formulations but if they are similar enough, the cells are usually be happy. We acclimate the cells to the L-15 for a couple days (in an incubator with no CO2) and use the same media during imaging.

The other thing to think about is where exactly the Oko Lab system is measuring the CO2 mix. If its measuring CO2 mix in the gas mixing unit, it’s likely that once the gas is dispersed into the air space of your microscope, it’s not 5% CO2 that the cells are actually seeing.


This is very helpful! Thank you for that. I’ll read up on the L-15 medium. Do you measure the CO2 levels close to your samples with a small gas analyzer or what would be the suggested way of doing that in a relatively easy/affordable way?

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We’ve seen both HEPES and L-15 work well with many cell types, but I’d recommend validating capability by growing your cells in the media in a tissue culture incubator for the typical duration of your imaging experiment.


I agree in Jennifer, very important to confirm growth with any of these mods. Re the 5% CO2, it should be 5% in the atmosphere of the chamber and local to your cells, not familiar with the chamber but many have a sensor in the chamber with feedback to the CO2 mixer. Are your macrophages proliferating? Playing with density to start with enough but not so much such that your medium is exhausted is a variable that is part of the art of long timelapse…you may have to explore perfusion if possible, adding medium during the run to account for what is lost due to the evaporation issue is not good as the osmolarity of the medium is changed due to evap, now you are adding more medium, but the osmolarity is still off…cells are very unhappy when this happens. Good luck with your experiments!


Thank you both for your suggestions @jennifer & @JamesOrth! The RAW 264.7 macrophages are dividing the first ~12 hours but stop shortly after (however… this may also be due to photo-toxicity). I try to seed them at around a 30%-40% density so that they have enough space to expand and that they’re not crowded. I will definitely check whether any of the modulations affect cell viability. I have been thinking about perfusion, but am still in the process of finding a good pump for that system. I had thought about adding medium, but also figured that the osmolarity would change. The chamber we have is the ‘cage incubator’: http://www.oko-lab.com/live-cell-imaging. I’ll try to find out where it measures its CO2 levels.

I am using the stage top incubator from Oko-Lab and able to do up to 10 days live imaging with no problem. Maybe, you could consider switching to such a solution or upgrade your set up. I’m sure Oko-lab will be happy to help you on that.

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@Maikel, how are you vizualising them now? Staining? Try to minimize exposure to excitation light. Both by decreasing exposure time during acquistion of each frame and by reducing the number of frames (increase interval). (I’m assuming you have a computer controlled shutter for your excitation light.)

Something to consider here: what do you really need to image to get your data of interest?
Do you need a frequent sampling rate or is total duration more important? Are you only interested in cell location or are you trying to track subcellular structures? Can you supplement your fluorescence images with phase contrast? E.g. if you want to track a subpopulation of cells that display a fluorescent marker, maybe you are ok with one fluorescent frame followed by multiple phase contrast images. One frame for identity, time resolved for tracking.


Good point! A few people i know bubble through a bicarbonate (?) solution instead of water to cut down on loss. There are 7% CO2 tanks or you could use a gas mixer, which is handy.


Thank you for you message @Henrik. We have a pretty pimped out Nikon Ti2, with Okolab cage incubator (CO2/Compressed air) and gas mixer. You’re right, we actually use camera triggered acquisition to minimize excess excitation. Our optics set-up can be found here: https://www.fpbase.org/microscope/qMBGEBUa9q9xe6k9VhfLGe/

I’m using a combination of dyes and fluorescent protein fusions. Ever since the CSHL course by @jennifer @talley and @Anna I have been far more aware of the choices you mention regarding intervals of acquiring images. I’ve also been avoiding BFP variants/near-UV excitable dyes because of photobleaching. Most of the imaging I do requires me to track individual bacteria through sub-cellular structures. I want to follow trafficking of these bacteria throughout macrophages, which requires a relatively high imaging interval. It’s interesting you mention using phase contrast as a tool. I’ve been thinking of using DIC or phase contrast to track bacteria within compartments between fluorescence acquisitions or even use some image analysis tools to be able to use DIC/phase information to track bacteria throughout the macrophage without using fluorescence.


We do 48-72 hr timelapses. Some quick general things from our experience:

  1. As others have said, measure CO2 within the chamber itself. Cheapest fastest way to do this with an IR CO2 meter (we use one from co2meter.com) and we calibrate CO2 levels within the stage-top incubator itself.

  2. Mineral oil can help with evaporation, but it can also cause depletion in oxygen from what I have read. See here: https://www.ncbi.nlm.nih.gov/pubmed/19672876

  3. Live-cell perfusion/media addition/removal is a great way to solve media evaporation. You can use a dual syringe pump with inlet/outlet needles, or a formal flow chamber, etc.

  4. Whenever possible, use DIC/Phase and image analysis/tracking. They are fantastic, low-toxicity, high contrast techniques for morphology, migration, and population dynamics. Of course they don’t always provide what you need, but when they do, they can dramatically simplify imaging.


Hmm. Bacteria within cells sounds tricky. I have no experience in this but sounds like you might need high temporal resolution and fluorescence for the bacteria. You can probably get away with unlabeled macrophages though.

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Some very top level things to noodle…

  • An autofocus (laser based) is almost a must-have, but as others have suggested being able to disable the IR laser or LED used for the AF function is also important.
  • It’s never a bad idea to test your hardware independently of the biology side via using a fixed test slide. At a minimum something in a similar sample container, to validate the computer, software, stage, microscope, etc etc. That way you don’t have to find out some random issue (like the computer deciding to go to sleep ) while losing samples at the same time. It’s also good because if things look reliable on the HW/fixed side it’s easier to diagnose problems if you have them only when running live samples.
  • A documented “check before clicking capture” protocol of some type can be helpful to maintain consistency, as it’s easy to forget how things were captured 3 days ago.