Hello all,
maybe I should have posted this in Long-term Live Cell Imaging tips/tricks but it would have been a little bit off-topic. I am currently doing live imaging on cells cultured in a 35 mm coverslip that I mount in an imaging chamber filled with medium. Because the chamber is not as sealed as a glass-bottomed dish, I am afraid that I might be killing my cells if I clean the bottom of the coverslip with 70% EtOH prior to imaging. What do you typically use?
Thanks!
Hi Irene,
I can’t say that I have any hard data on this… I haven’t ever worried much about it. It seems like a cotton-tipped swab and a little bit of care not to get close to the edges of the chamber should be enough, even if you use ethanol. I guess it also depends on what you’re trying to clean off. If it’s just dried culture media or other salts, then even just water or diluted glass cleaner might be enough… if you’re cleaning oil then you’ll probably need a stronger solvent (or you can just decide not to reuse a coverslip once oiled).
If you’re really worried, i suppose the safest approach would be to just do the experiment one day: clean some, and don’t clean others and see if there’s a difference.
Are you cleaning the coverslip in order to sterilize it after being submerged in media or because there are salt/media/etc adhering to the glass and interfering with imaging?
I used 35mm coverslips that had been grown up in 6 well plates extensively. We had a metal holder we would install it into and then used a cotton swab with a small amount of ethanol to clean the bottom of the coverslip (the slides were infected with Chlamydia, so we didn’t want to rub that on an immersion objective.
We never had any problem with it affecting cells (there were still going through the cell cycle just fine for 48 hour experiments), so as long as you have the cover slip installed into a holder with a tight gasket there should be no issues.
(what we used, but anything similar will work-- https://www.thermofisher.com/order/catalog/product/A7816)
Thank you all for your replies!
@JawnnyH, I typically do so to prevent imaging interference from the salt adhered to the glass. I could try with water, as @talley suggested. Unlike @H_Brown-Harding, I don’t have a potential problem of contaminating the working space with a pathogen. Our chamber is similar to the one posted by Heather but the closing mechanism is a clasp. There could be a handful of reasons why my cells are unhealthy so I am checking everything to the tiniest detail.
@talley, right now I am also removing the immersion oil after imaging so I can continue growing the imaged cells for a little longer. So far I have used 70% EtOH and it’s not the cleanest. Do you think using chloroform and a cotton swab to get rid of the traces is too risky for the cells?
If your main concern is salt, I would just use water like @talley said. If the salt came from media, it should be the most soluble in aqueous solution and if you’re only use 70% EtOh, you might cause certain salts to precipitate more.
There are many wonderful protocols out there to clean slips for cell culture, I have used several, some which include acid stripping, sonicating, and autoclaving. You are right about residual ethanoll, the tiniest amount will be toxic to your cells (no liver :-). A method I settled on that allows for bulk preparation is: 1) put a standard pack of slips into a 500 ml beaker, 2) wash 2-3 times with excess 70% ethanol, 3) rinse 2 times with 95 or 100% ethanol (this removes water and helps reduce the slips sticking to one another), 4) place foil over beaker and autoclave. This way you can prep 100 slips at once. As long as you open the beaker only in the laminar flow hood, you can use the entire set of slips without incident. Some people simply store slips in 70% ethanol and then take them out with forceps and flame them, the put them into the dish. If you flame too long the slip with splinter into pieces and this is time consuming.
One question I have though, are you placing a glass slip into a plastic dish (with cells on the slip), and imaging through the plastic via an inverted? This could be ‘ok’ for brightfield, but will diminish fluorescent performance significantly.
Glass bottom dishes are much cheaper than they used to be (not MatTek), or you can make your own by drilling a hole in the bottom of the 35mm dish and using epoxy or super glue to put on your own slip (did this in grad school) - dirty little secret - you can even reuse them!
Good luck!
I think I misunderstood your question. To clean the bottom of the slip, I’ve used isopropanol or optical cleaner (or sparkle). I’ve never had an issue cleaning the bottom of the slip.
Good luck.
We typically use a tissue to remove debris and medium from the bottom of the coverslip after mounting in an imaging chamber (we image on an inverted microscope and removig the medium from the coverslip bottom is important to avoid mixing of medium with immersion oil).
I recommend cleaning with a kimwipe damp with distilled water, then wipe dry.
Do not forget to clean those Attofluor-type chambers as well! When doing pharmacological studies, we found that following experiments would be impacted (to varying degrees) unless we cleaned the chamber aggressively between uses. I generally pop out the gasket with a razor, wipe all parts clean, soak it all in 10% acetic acid for 30m and then wash thoroughly with dH2O.