If I could get some help here I would be very very grateful. I am having such massive problems with cell culture that it is seriously affecting the progress of my PhD, and it's starting to get very depressing as nobody seems to have any answers for me.
I was using a cell line called 10T1/2 which is mesenchymal and grows in MEM supplemented with NEAA, 10% FBS, 2mM L-G and P/S. I cultured this line for about 6 weeks, passaging was fine the cells seemed reasonably easy to look after. When I froze down aliquots (LN2) and thawed them out I had no problems. All is well. Then suddenly it wasn't- the cells would not revive anymore, they would lay floating in the flask (this from batches which had previously worked) I obviously made new media up, cleaned the incubator, checked the temp (I cant check the CO2, other than believe what it says) but I used another one in the lab etc. Nothing made any difference. No one could tell me what I was doing wrong.
I couldn't get hold of any more of these cells, so I switched to another line ATDC5 cells. We have a limited no of aliquots a previous lab member froze down and had done a lot of work with. I thawed these cells out, standard procedure 2 mins in a water bath, ethanol spray, into the hood, span 1 vial down (appropriate speed), the other I just seeded right away- T25 flasks, with warm complete media. The media I use is as stated on ATCC, UK cell repository etc etc DMEM/HAMSF12 1:1 mix, supplemented with 5% FBS and P/S mix.
I seeded the cells at 11am, I checked the cells at 5pm the same day they had attached and had taken on the characteristic fibroblast like apperance, the media looked good in colour and appearance. The next morning at 9am I checked the cells under the microscope (I checked each flask individually, each one was out for less than 1 minute) they looked great, fibroblast like attached and as though they had grown well- no problems so far. The media was also normal pH.
After 24hours you are supposed to give cells fresh media after thawing, so I planned to do that. However at 10:30am that day when I checked them again before changing the media- the cells had rounded up and become detached, completely useless to me and most definitely dying. Literally one hour previously they had looked fine.
By 1pm that afternoon you can see cell debris and all cells are rounded and floating.
The only things I can think of are:
Mycoplasma ( I have streaked some culture onto an agar plate to test ) although this does not explain why at 9:00am the cells were fine and by 10:30 am they were not. Also I dont believe they would outright kill the cells.
Incubator: Temp is at 37 (thermometer) cannot check CO2 but readout says 5%. Also cells were left in this incubator O/N and were fine, they were taken out for less than a minute to check under the microscope (very gently i will add) and then placed back in.
Media: The FBS we buy is quite cheap I believe compared to some other products (£30 for 500mL from our University stores) it is cell culture sterile and filtered, FBS from south america. Other cell lines have been cultured using this, but perhaps this cell line doesn't like it? However I would imagine if this was a problem- the cells would not have attached and grown O/N in the first place?? Why would they just randomly die like they have done??
Please someone help me!!! I cannot think of a single logical explanation for the cells being ok at 9:00am the day after seeding and then not being ok at 10:30 am the same day. I don't have many aliquots so I can't just keep randomly trying. Plus with my absolute failure with the previous cell line, I am starting to think there is something wrong with me/something I am doing.
Thanks very much,