Why, why, why? can I not keep cells on glass?!
I (anybody) can get them to stay on plastic, no problem. And they’re fibroblasts – bloody self-spot-welding limpets! Stick like shite to a blanket! But, trying again on glass coverslips (because the plastic supposed wonder-product for fluorescent microscopy coverslips turn out to be useless for bright-field), and I’m reminded how haphazard keeping them on glass is – or how inefficient I am. Despite all(?) precautions: washed in detergent; rinsed repeatedly in distilled water; immersed in acid-alcohol for one hour; air dry in the hood. Then coated overnight with fibronectin and gelatin; then plate the cells. And they seemingly attach fine; they look the same as on plastic. It’s when I come to fix them. They come off in a sheet (which I suppose suggests they hadn’t attached properly at all). Despite my carefully applied render, they’re rendered uninformative. Just (a) floating gossamer of gormless cells.
Sometimes it works; other times, like this morning, they detach so readily, that others in the corridor are commenting on that apparently Tourette’s-afflicted bloke in the lab who has just wasted another two days in an effort to nail a (probably pointless) figure.
Hang on, I’ve just thought of something…
I had this same problem both with peripheral neurons (ephemeral fly-by-night things) and with peripheral nerve sheath tumor cell lines (more limpet-like than fibroblasts) made from my knockout mice. The neurons required a polyornithine and laminin coating even on plastic, but the PNST cells did not. I wanted to image mitochondria on the confocal, and I couldn’t get the cells to stay attached. Finally, in frustration, I made a strong HCl solution and left the coverslips in it for an hour or so. After several water washes, I soaked the coverslips in EtOH, and then washed them again in autoclaved water. Some were quite brittle and broke, but the intact ones were propped in petri dishes to dry in the TC hood.
My hypothesis is that the strong HCl etches the coverslip glass somehow, leading to better attachment, but I have no evidence in support. A research assistant in a neighboring lab also used this approach with success, and I think she actually measured how much HCl I put in 100ml of water, in case you want to try this method. I tend to relinquish my OCD when it interferes with experimental progress.
Transfect the cells with talin. That’ll make the bastards stick, for sure.
I also suggest a polyornithine pre-coat before putting ECM proteins on. In my day, we started also with poly-D-lysine, if I remember right. We did that for neurons on plastic as well, mostly because it was a ready protocol. This is not an endorsement, but just to give you an idea.
I also remember being told to not bother cleaning the coverslips, as if they are too clean, the cells can slide off… :-) YMMV.
Wow, I never bothered to clean coverslips that thoroughly. Couldn’t be bothered. Collagen coating always worked for me. I found that overconfluent cells tend to detach more easily, coming off in a sheet. Particularly if the fixative is very cold. Perhaps fewer cells and warmer fixative would do the trick?
I absolutely hate it when minutiae hold up my work. If it’s something big that requires thought and reasoning to solve, fine. If it’s something dumb and routine… oh, how I hate that.
Thanks for the tips. I slept badly last night, went into work with eyes like onions, then couldn’t get on with it properly because the hoods were being tested (which I’d been alerted to a few days ago, but forgot). So, the last thing I needed was the cells taking the piss. Hence, in a fit of pique, I decided to rattle off a post about it. Don’t know why, but it made me feel better. (Does anybody else get that?)
I once read/heard that glass coverslips can have a film on them, and detergent and acid gets rid of that, so cells don’t ‘slip’ off. And, of course, ethanol to sterilise. Never occurred to me that they could be too clean. And I’m not sure whether it’s the coating not taking properly, because that’s what the cells need; so if that’s loose, the cells won’t easily hang on to bare glass?? (Or do they – acid-etching? Seems feasible.) I’ve only used poly-L-lysine to coat slides for cytospin purposes, which works fine when you’re not bothered about morphology (having taken them off the culture surface).
But I think I’ve twigged why? I’d plated with serum, which contains factors that aid attachment. I find I need to do this when plating these cells on plastic also, else they don’t attach properly. Once attached, however, it is okay to switch to serum replacement. But, not on glass. And the annoying thing is, I didn’t need to switch, because I wasn’t plating other cells (which I culture in serum replacement) onto the fibroblasts (feeders) – because I just want to test the feeders! It has worked okay before, but that’s because I kept them with serum. I think. Something to test, anyhow (when I’m in the mood).
Along those lines, keep some Ca++/Mg++ in your buffers, if you don’t already.
Good one! I bought in some PBS with them way back. However (is that the sound of a penny dropping?)… I didn’t make up the PFA !?!?!?!?!
_It’s when I come to fix them. They come off in a sheet _
YES. This happened to me countless times when I was using chamber slides (which are also glass, but pre-coated, so I never had to do anything to make the cells stick).
This is how I solved it:
-Don’t grow them too confluent. That’s what’s making them come off in a sheet. If you seed fewer cells, and they don’t lift off, you’re eventually going to have more cells than when they’re confluent and all jump off together.
-Don’t shake anything, ever, at all, while washing/fixing/staining. Not even after fixing. Just don’t. Don’t move the slides/plates. Don’t bump into the bench they’re on. Just very carefully add/remove the different liquids.
I still go things lifting off around the edges of the well, but that was in chamber slides. On coverslips in big wells that shouldn’t happen at all.
PFA must be fresh (reduces autofluorescence, too). Also stick in ~2% sucrose.
Or freshly thawed… but I’ve never bothered to put divalent ions in my fix. Sucrose is helpful for maintaining nice morphology, and can’t hurt. And the PFA must also be buffered, as it’s naturally quite acidic – I remember a whole series of messed-up experiments from that one, using a stock someone froze without checking the pH.
I like your turn of phrase concerning a “floating gossamer of gormless cells” – those of us who have seen it, know exactly what you are describing.
These particular cells are irradiated, so don’t grow too confluent. And, it has, in the past, sometimes been okay. (Intermittent snags – always the worst, because you think you’ve sorted it, when you haven’t).
Yeah, ‘we’ prepare and freeze aliquots of fresh PFA prepared in PBS, and thaw when needed, and discard that not used within a few days. But, what PBS is it prepared in…. (am on the case)? However, I’m pretty sure they were loose anyway – the serum / replacement thing? That’s my working ‘hypothesis’ anyway (will let yous know).
‘Frozen’ != ‘fresh’
(although that doesn’t sound like your problem, anyway.)
No, that doesn’t matter. I always used either fresh PFA or week-old PFA (never anything that was opened longer ago) and between the two there was no difference in non-fixiness.
Sorry, I cannot help. But feel free to contact me when you need advice on how to kill cells. At that I am expert.
Oh, killing cells is my inadvertent forte. In fact, maybe that’s my response to the party question… “I’m a cytocide.”
I’m talking about AF, which is different from non-fixiness. Which I admit wasn’t the topic but that’s never stopped us before.
Okay, if I can diverge into autofluorescence, I’ll bite. Why would older/fresh-frozen-thawed paraformaldehyde give more AF signal than freshly-prepared? Yes, there may be less polymerized formaldehyde floating around, but what if you treat with sodium borohydride (to reduce any available aldehyde groups) or glycine? I thought it was free aldehydes that sort of non-specifically fixed your secondary to the tissue. And has anyone anywhere done any sort of kinetics on the polymerization and how that relates to optimal (read, low background) fixation, or is this all lab lore?
There’s a difference between AF and non-specific fluorescence, which is what you’re talking about. I understood that it was long-chain PFA that gave the signal.
There are some good tips about “AF” assembled here, by the way, found through Internet trawling.
Quite right, Richard, I was conflating.
I use neutral buffered formalin – always prepared fresh – for fixing tissue, since getting some nice TUNEL results. However, I’ve not used it for cultured cells, which only take minutes to fix. PFA depolymerises in solution; I wouldn’t have thought repolymerisation should be a problem, providing it’s ‘fresh’ enough.