Dear All,
In my experiment, I need cell to be 50% confluent in 100mm dish to treat drug and collect samples after 24h.
To prepare cell test group (I use large number of cells. 8 of 100mmdish), Which is better between:
(1) subculture cell from the same one flask into mutiple vessel equally, and let grow to wanted 50% confluent, then treat the cells.
(2) subculture cell from the same one flask to multiple flask first. Let all flask grow, then collect all cells mixing together into single solution, count cells, then subculture 50% confluent cell number into each new dish, equally. Then after 24h (lag phase, no cell number change) treat the cells.
The idea of first method is for eaiser to distribute small cell number for preparing many group of large number of cell, but not sure whether cell grow/become more difference (in cell number between each group) in longer time. While, the idea of 2nd method is for control cell number, because the cell is in lag phase (after passage 24hr) and less likely to grow more from what it has been passaged.
Should these two method be the same?
Thank you
if you have more number of cells, try to use in initial passages especially for endothelial cells like HUVEC, in case of highly dividing cancer cells two or three passage does not make any significant differences.
Dear all,
when cells reach confulency, signal transduction pathways for stress may become activated. So you may have false positive results.
So the best way is to treat cells with the cytotoxci drug, when they reach 60-80% confluency, depending on the treatment.
I think the right path needs to be empirically determined- try each way and evaluate performance both in terms of altered signaling in response to the drug treatment and for reproducibility (direction and magnitude). Any of the following variables might influence your results: passage #, size of culture vessel, length of time between cell seeding and drug treatment, cell density/confluency.
If cell density matters in your experimental system (worth testing directly), the two methods you outline will not necessarily give the same results. Certain cell types tend to grow non-uniformly, leading to variable levels of confluency across your culture vessel. On the other hand, seeding cells at high density can lead to high concentrations of cells in the center of your well, and sparser at the edges, especially in small culture formats (12 well or smaller). I would NOT expect to see non-uniform seeding in 10cm (or larger) dishes provided you evenly disperse the cells when initially seeding the plate.
Whichever method you choose, make sure you have biological replicates (n>=3) from the same day. Otherwise, it will be difficult to assess which experimental set up is better.
What type of cells are you using?
Thank you all.
Dear Daniel, I use SH-SY5Y cell line. I already repeated the experiment many time in six well and find out at cell confluence around 80% give best signal response to the drug. However, I need to do it in 100mm petri dish in my next experiment. I normally culture cells in T75 flask, which is not enough to distribute confluent cell number into 6-8 of 100mm-dishs.
I use the 2nd approach, harvest near confluent cells and seed into different dishes at equal and appropriate density for the dish, incubate in complete medium overnight, remove medium and incubate in medium without PBS for at least 1 hr before treatment. For my cell lines, the treated cells are at exponential growth phase. Since I use untreated cells under the same culture condition as control, I am able to do comparative studies. The 2nd approach would seem to correspond more closely to the in vivo condition at which the drug mediates its effect.
Agreed, you probably need a T75 flask of cells for each 10cm dish seeded at confluency.
The second method seems more controlled, both for number of cells and for amount of nutrients in the medium (since that was changed more recently).
Hi, I have to agree with Daniel and many above. It depends on your assay. However, in general you should use the same passage of cells within one experiment, therefore your second method is more controlled, you can synchronise your cells, would give you more cells at the same passage and in general you should use confluence 70- 80%. In addition you need to keep in mind that actively dividing cells are more susceptible to treatments.
If you absolutely MUST treat at 90%+ confluency, then seed the cells the night before your experiment at a density that will yield 90% confluency in order to minimize the influence of intercellular signalling. If you let the cells grow to this confluency vs plate to this confluency, your results will likely be more obscured by intercellular signals. Keep in mind that your variability will likely be higher than if you perform the experiment at
That number will be different for every lab. Others working with the same cell line are subject to variables such as different lots of FBS, treated vs non-treated culture dishes and if antibiotics such as pen/strep are used. It is best to run a small seeding experiment to determine that number.
It depends on the rationale of your experiment, the type of drug you are testing , and the effect you are trying to measure. If you go with #1 it is normally advisable not to let the cells grow beyond 70-75%. so you can see the effect on certain cell signaling pathways. It depends what you are interested in. At this point the majority of the cells would have cell-cell and cell-ECM connections established, thus you will not get signaling that usually gets turned on when you lift the cells. Since there is crosstalk between these and proliferative and survival pathways, you may get skewed results, unless you account for these or purposely decide to test cells at this point (option #2).
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