No. Indeed one cannot use any assay that is based on adherent cells becoming labeled. If 'X' causes an increase in ROS, hence Mitosox signal and cell death, the the amount of signal per cell increases and the number of cells goes down.

You have to fix the cells and use flourescence microscopy to determine signal/cell and go from there.

Thank you, Rodrigo. Martyn, I understand that cell death would change the number of cells. However, if I have determined cell survival for that particular cause "X", then I know how much it contributes towards it. And if assay is performed within 15-20 minutes, after the treatment, will that really change the signal ?? Can you please explain that argument again ?? Thanks !!

Hi Rashmi, you can do a kinetic assay in a fluorometer. We've been using this method in our lab with macrophages and it works well. It is essential to use mediums free from phenol red. Another tip is to load the cells with the dye in the absence of serum. I think a good idea is to use MitoTEMPO, a mitochondria-targeted antioxidant with superoxide scavenging properties. This will help you to show the specificity of MitoSOX.

Good luck!

Thank you, Fabianno ! Even I am planning to do kinetic analysis in myotubes. Is it possible for you to attach reference for this sort kinetic analysis for mitochondrial ROS ?

Hi Rashmi,

I think what Martyn is highlighting is the mode of action of MitoSOX. Sure it interacts with SO, but the oxidation product does not become highly florescent until binding to nucleic acids. So with cell death comes an increase in free nucleic acids and SO in your culture and thus presenting a false positive. Even with your accounting for cell death can you account for false positive defined as increased emission of signal?

The question could be answered if you have not found an increase cell death during optimization of your treatments and a mitochondrial targeted SO scavenger, as Fabianno suggested, produces a marked reduction in your signal when used in conjunction with your MitoSOX assay. I hope this helps.

Good luck!

Thanks Micheal !! I will keep these things in mind while designing the experiments.

Rashmi , I understand that cell death would change the number of cells. However, if I have determined cell survival for that particular cause "X", then I know how much it contributes towards it. Can you please explain that argument again ?? Thanks !!"

Here is the problem. You have mitochondrial signal and a small background signal from the proteins sitting at the bottom of the plate. Each control cell gives you a signall of 100%, background give 5% and you have 70% of the plate surface area covered with cells.

Now you add 'X' to half the cells and at time t add MitoSox. Now anything that is going to increase mitochondrial superoxide generation is going to be cytotoxic. The more ROS generated, the more cell death.

Imagine if treated cells give a signal of 600%, but dead cells give 0%. Small changes in cell numbers convert a signal of 600% into lesss than background. You get titrations that begin at 100%, rise and then fall. The ROS levels rise, but total cell numbers fall.

You cannot use Hochst or other viability dyes to count cells and normalize, as dead/dying cells tend to be brighter.

I have another query with regard to use of MitoSox Red in 96 wells plate assay format. How do you find optimal concentration of Mito ? Is microscopy important before going for 96 wells plate assay ? Because I think determining specificity is important. However, another method seems to be using Mitotracker. So can anybody help me out with this, may be by sharing their experiences ??

Dear Rashmi, have you been able to optimize your experiment on 96 well plate?

I am currently struggling with similar experiment: I treat the adherent cells with 5uM Mitosox in normal media (including phenol red) for 5 min in dark (foiled), 37C incubator. Then, I wash cells 2x with PBS (as instructed from the manual). Then I add PBS and take it to the plate reader. To read fluorescence, I use 5nm step and sum all the fluorescence from 565nm to 710nm. I normalized the signal by the protein concentration from the cell lysate (from the same plate). Mitosox is diluted in DMSO from power just before using. 

I am wondering if my conditions are okay. It takes too long to measure (perhaps 5nm step size is too narrow?). Do you suggest that I incubate cells with phenol red free media and also read the signal in the phenol red free media (with serum?)?

Thank you!

Hi Eunjoo,

Yes.. please use DMEM without glucose without phenol red for incubation as well as for washing instead of PBS. It will stabilise the signal. Mitosox is dissolved in DMSO. For reading the signal, remove the medium completely and take the reading. It would be a good idea to measure total protein from the well or total number of cells if possible to normalise the per well flurescence. Hope it helps. All the best !

Rashmi

Thanks very much! Rashmi I used a media without phenol red for incubation and washing. I also removed the media completely as you recommended. Then I read the fluorescence by a microplate reader. Then I normalized the reading by protein concentration. This gave me a great signal. Thanks very much!

Dear Eunjoo,

How are you performing your BCA assay for protein quantification? Are you carrying out the whole protocol in the original 96-well plate? if so, how much RIPA buffer are you adding to each well?

cheers.