Hi Suleyman

It depends on how you want to plan your experiment. Your control has to be in the same conditions as your treated samples. Meaning, if you incubate your treated samples for 12 hours, your control should be incubated for also 12 hours. If you do a media change (for example) to refresh your drug, your control also has to be refreshed (but only media, not drug).

I have worked in the past with kinetics experiments with suspension cells. I wanted to see the expression of a specific protein at 4, 6 and 8 hours being stimulated with the drug. Since this is a very time-restricted experiment, what I did was to seed one well for my control (unstimulated), and one well for every hour or time point I was analysing. Then, I was treating my samples in a backwards sense. For example, lets say that at 10 am i add the drug to my 8 hours well. Then, at 12 noon i will add the drug to my 6 hours well. Next, at 2pm i will add the drug to my 4 hours well. And finally, at 6pm my experiment will be finished, as the longest time point (8 hours) is complete. Hence, you collect/lyse/whatever all your samples at the same time. That way, because your collected everything at 6pm, your samples have been treated the time you wanted (meaning, the 4 hours time point has indeed been treated 4 hours because you added the drug at 2pm and you lysed the cells at 6pm). And your control has been incubated in the same conditions as your longest incubation , which is 8 hours. This way you only have to add one control.

Now, I have had supervisors that disagree with this, as technically, you do not have one control well for every time point. Currently, I am doing treatment experiments in adherent cells, but I am looking into weeks long treatments. So in this case, I have one control well per time point. My first time point now is 24 hours, so 24 hours before the treatment, I treat all my samples. And then I collect my cells at 24 hours, 3 days, etc, after treatment, including the control well of every time point. Note that in this experiment, for example for western blot, every treated sample will be normalised to the control of that time point. But in the other experiment i explained above, you will normalise all the treated samples to a single control.

if you are looking into hours long experiments, in which the maximum incubation you have will be one day, i would suggest to do the experiment in a backwards sense. Your control should not be changing that much as it is a short period of time to investigate. Contrarily, if you want to investigate longer time points, i would add one control per time point. You can also do the experiment backwards, but still adding one control per time point. Your cells can change while being plated on the tissue culture plastic, specially if they grow because of the serum, or you do media changes, etc. In that case, it is better to add one control per time point

i hope this makes sense!!

ps - if your drug is dissolved in something like DMSO, i would suggest to add an extra well of this condition. So you treat that well with your DMSO concentration, in order to see if this stimulates the protein you want to see. (likely not)

good luck!

Hello Noelia,

Thank you very much for these long explanation. It is very helpful indeed. Generally I am giving drug diluted with complete media and always I add 10% of total volume. Like if I make total volume 5 ml, 500ul becomes my diluted drug at desired concentration (by the way I hope I am not making mistake here). For control I am adding 500ul complete media (drug is very high concentrated instead of adding 1ul DMSO per 2ml, I don't add). In your case, for example I plates 4 tissues (cont 2h 4h 8h). 5ml drug and medium will be for cont and 8h sample and 4.5ml for 2h and 4h samples. After 4h and 6h later I will add 500ul drug. This is correct way to use it right? Besides for adherent cells, should I use the same approach?

Thanks in advance.

süleyman

Hi Suleyman

I am glad you find it useful. Your calculations about the 10% of total volume are correct. I assume in those 500 uL you add to your wells your drug is in an specific molarity. It is always good to add the drug already diluted in media, that way you do not make mistakes pipeting (specially if you need to add small quantities). However, I always used to add the drug in the total volume of all my wells. Meaning, if I have 4 wells with 1 ml each, that gives you 4 ml of total volume. If you multiply how much drug you need per well * 4 total wells, that concentration you dilute it in 4 ml of media. And the day of your treatment you remove the entire volume and replenish with 1 ml/well. This is quite efficient with adherent cells, but for suspension cells, the way you described should be okay, as likely your cells will sink to the bottom of the well and it is easier to add 500 ul of your diluted drug, as long as you end up with the concentration of drug you desire. And the way you describe how to add in the different time points it is okay.

For adherent cells, you can use the same approach. Just bear in mind, if you want to do western blots or maybe RNA isolation for PCR, you will need to lyse your cells from the well, or detach them with trypsin and then lyse them. If you do this, I will recommend you to use one plate per time point. If you have all your time points in the same plate, you will likely spend some time lysing or detaching your cells, and the rest of your time points will be outside the incubator etc, and in a restricted time window, that can affect (in my opinion). Specially if you want to lyse your cells directly on the plate, as that will take some time. Also, if you ever use trizol for RNA isolation, that contains phenols that are highly volatile (even more at 37 degrees) so if you have all your time points in one plate, and you lyse one well with trizol, and put the plate back in the incubator, the rest of your wells can get contaminated and hence die! For suspension cells it is easier as you just pipette the cells and continue working.

Hope this is useful as well!