Discover the world's scientific knowledge
With 160+ million publication pages, 25+ million researchers and 1+ million questions, this is where everyone can access science
You can use AND, OR, NOT, "" and () to specify your search.
I am trying to perform a MTT assay, and I have found some unexplained problems.
The protocol that we are using is the following:
1st day: Cells are seeded in 96 well plates (5000 cells per well) and incubated 24 hours.
2nd day: Wells are washed with PBS, and different concentrations of the materials are added, together with medium alone as control.
3th day: After a 24 hour incubation, medium with materials is retired, MTT is added (0.5 mg/mL per well) and plates are incubated for 3 hours. After that, MTT is retired, and DMSO is added to solubilize the formazan crystals.
When performing the experiment, we found that in some wells, apparently at random, cells in the centre of the well began to degrade, appearing with a round shape, and even membrane blebbing in some cases, while cells in the periphery remain healthy. It happened in different steps of the protocol: Sometimes, after adding the materials, other times in a moment between the addition of MTT and the addition of DMSO. This is particularly apparent when we check the cells after adding MTT, since you can see that the formation of formazan crystals is totally inexistent in the centre of the well. It looks as a toxic compound dropped in the centre of the well.
We tried changing the plates, and the result was the same: In some of the wells, this problem appeared. The cell line is growing well in the flasks, and after the first 24 hour incubation in the 96 well plates they always look healthy. This problem only appears after manipulating the plate.
I have worked before with this cell line, performing similar protocols, and I have never had this problem. Anyone can help me or give me any advice?
Thank you in advance.
I've recently started working with human aortic endothelial cells. I acquired a vial from Lonza, and I feed these cells with their EGM-2 medium which comes with the many growth factors that these cells require. Up to the first 3 passages, everything was going well. However, the 4th and 5th passages look terrible: there are hardly any adherent cells, and there is a lot of dead cell material in there such that the medium looks pale pink and chunky/turbid. I have a few freezer stocks of these cells, but I am reluctant to thaw these until I know what the problem could be.
When I split these cells, I use accutase (instead of EDTA or the harsher trypsin). I haven't changed cell culture flask brands either (Corning T-75s throughout).
I'd appreciate any input/ideas. Thanks!
I am working with primary HUVEC cultures from Lonza. They are stored in liquid nitrogen at P5 in a freezing media of 90% FBS and 10% DMSO. They are plated on TC dishes coated with fibronectin in VascuLife EC media with VEGF at a density that has always been successful in the past. Media is changed the day after plating, and then every other day thereafter.
Sometimes the cells are able to proliferate and form a monolayer normally. However, in the past month, many of them experience death in large chunks of the plate starting the second day after plating (so after the first media change). For example, in a 6 well plate, one well may proliferate normally, two may experience almost complete cell death, and three will look normal in >50% of the well, but gone in the rest.
What is really confusing me is that they "balloon" out before death, a process that I am not familiar with. This occurs in cells handled by all lab members, so it is not one individual's technique in particular.
I have attached a 20x image showing a small area of healthy cells next to the sick ones.
Any guidance on how to remedy this issue would be greatly appreciated.
I have worked in NCI-H1650 (human non-small cell lung cancer cells), I have thawed these cells for 3 times, in all the cells were ok and attached well after 24hr, but after 48hr the cell is dying and all deattached, I have applied the same protocol as in the literature, Dose anyone have any suggestion about this problem?
I am working with primary human lung fibroblasts and when I go to either treat the cells or transfect them, they die (sometimes). The cells are maintained at 37degrees in a humidified incubator in an atmosphere of 5% CO2, and fresh growth medium are added to the cells every 3 days until ready for experimentation. The cells are maintained in the same incubator, but a serum free medium for 24hours before the experiments.
I treated the cells with TGF-beta at 10ng/ml in a 12-well-plate, and seeded 200,000 cells in the wells. when i checked the cells the following day, a lot of them had died. Why is the the case because, sometimes they die, other times they are fine.
I've been having this problem for the past week now and I don't know what to do anymore. Following siRNA transfection a week ago, all cells I plated on coverslips died. I felt it was because I used a high volume of transfection reagent (But that has never been the case). So, I decided to seed a new batch onto coverslips and the same problem occurred after 24 hours. Thus, I decided to recover a new vial of low passage cells from liquid nitrogen. All cells behaved well and I went ahead to seed them onto coverslips once again. However, following siRNA transfection 24 hours ago, the cells underwent massive death once again. I'm totally confused as I've never encountered this before. What could I have done wrong.
I am trying to establish stable cell lines expressing my transgene. The transfections seemed to have worked fine and I had picked the clones few weeks back which were first growing in 96-well plates. I have been expanding them and now that I am transferring them to 12 or 6- well plates, even 24-well plates, they are forming clumps and are dying. They are not adhering to the plate surface anymore. Everything has detached, forming clumps which are swimming around. What could be the reason.
Hi all, using a transwell assay to test migratory function of cytokines, Costar transwells 5 micrometer, 24 well, according to publications and protocols 100 ul in top compartment with 1x10*6 cells (huPBMCs)/ml and bottom with 600 ul media10%FCS +/- cytokine.
Time titration 2-4 hr, and all I see in cytokine as well as in the media only control is cell debris.
Any advice? Very appreciated. Cheers, Axel
My CHOcells are not growing anymore and started dying after splitting for two times. I added fresh medium(alpha-men) with new FBS, and I still see the same result. If stocks are contaminated why they grow very well for first two rounds?
Our lab recently did a test run mixed lymphocyte reaction (MLR) using normal human PBMCs.
Medium composition was HyClone RPMI-1640 +10%hiFBS +P/S +Fungizone +2.05 mM L-glutamine.
The feeder cells were treated with mitomycin C at 50ug/ml for 1 hour to render them non-proliferative.
DPBS was put in the space between wells to help minimize effect of evaporation and edge effects.
Cells were grown for 6 days at 5% CO2/100%RH/37'C. Incubator logs show no deviations.
Excess stock from which cells were taken was plated in a six-well plate in the same media and grown alongside the MLR plate. These cells are still alive. The 96-well plates used for MLR are brand-new. There was no sign of any bacterial, fungal or yeast contamination.
MLR is new to me, so it may be that I'm missing something obvious. What's going on here? Why are all my cells dead in the MLR but not in my 6-well plate?