ROS measurement - Can someone help me? (Oct/18/2009 )

Hi.

I want to measure the ROS production caused by the treatment of some drugs in a cell line. I have two probes to do this: dyhydrorhodamine and dyhydroethidium. However, what I thought would be an easy task to plan, is now becoming confusing. Can someone advise me a good protocol?

Usually I do 24h treatment periods for some other assays (cell cycle, mitochondrial membrane potential) and I would like to mantain this condition for ROS determination. However, if ROS production occurs in the earlier hours of the treatment, will I be able to detect ROS after 24h? Should I do time-points determinations?

Also, the time for the addition of the probe is confusing me. What is more correct: incubate the probe just before the assay or combine it with my drugs at the beggining of the treatment?

I thought that I had to use flow cytometry to analyse the data. But, I'm seeing some protocols where the measurement is made with a spectrophotometer. Is this correct?

Thanks for any responses.

When I measure ROS, I first treat the cells for whatever time I'm interested in (24 hours in your case). Then I wash the cells x2 and stain them in FACS tubes with DCF-DA in PBS or media for 15' at 37 degrees in the dark. I then wash them once with PBS and analyze immediate by FACS. I'm not sure about the other probes you mentioned. I've never used those, but I've had good luck with DCF-DA. In my case, I've definitely needed to do a time course experiment to determine when ROS levels change in the cell in response to the drug treatment. My biological readout is generally death, and I've found that ROS levels change well before cell death occurs. Good luck!

Thank you for your answer.

I'm using adherent cells so I have to trypsinize them. I read somewhere that trypsine could distort the ROS measurement. Is it ok to incubate the cells with the probes before the trypsinization?

Thanks again.

Trypsinization increases ROS so it will give you inaccurate measurements.

Cells do not like to have DCF inside, they (live cells) will excrete DCF back out into the media. So incubate with DCF after treatment and only before you're ready to assay. If you plan on using DCF, buy it from Molecular Probes because the DCF from Sigma leaks like crap.

DCF is a fluorescent dye, you can assay with anything that measures fluorescence... the reason why people use different equipment is because it's cheaper to use which ever tool that's readily available.

Thanks.

Other responses are also welcomed.

I still have one question.

If I do the probe incubation at the end of the treatment period, ie, after 24h, will I be able to detect ROS formation that occurred in the first hours of treatment? Or will I obtain subestimated measurements?

No.

You will have to do separate experiments with different treatment times.

i.e.
EXP1 treat for 1hr, then do DCF assay
EXP2 treat for 24hrs, then do DCF assay
compare EXP1 to EXP2 to see if ROS occured during 1st hour.
(also include other controls: untreated/empty vector/etc)

cardosopedro on Oct 19 2009, 12:05 PM said:

Hi, like Bill Nye said before do separate experiments.

About your probes: dihydoethidine is a specific probe to measure superoxides, while rhodamine is specifi for mitocondrial membrane depolarization.

If you will measure the superoxide production for the 1st hour after the cell treatment you can incubate the fluorescent dye together an see it using either FACS, fluorescence microscopy or fluorimeter.

If you will measure for a long periods as for 24 h you have to wash your cells with PBS and then incubate the cells with the dihydroethidin in the dark and under agitation for 30 min. You can cover your samples with tin foil.

But if you want to measure the total ROS formation use the DCF dye as suggested because it is inespecific reacts with all kinds of ROS.

If you want too measure hydrogen peroxide the most recommend is to use the amplex red.

Thanks.

Hello Miss Montana,

I am using H2DCFDA (Invitrogen, Cat. No. D-399) for detecting intracellular ROS invitro. We are measuring the fluorescence on a multiplate reader. I am having some problems with this assay. can you please help me in solving these issues ?

In brief my protocol is:
1) Seeded 5x10^4 HepG2 cells/well in a 24 well plate, and incubated for 48 hours
2) washed cells twice with PBS
3) treated cells with 1000uM, 100uM & 10uM H2O2 and also included a non treated control. Incubated for 2H
4) washed cells twice with PBS
5) replaced media with 10uM DCFDA in PBS in a dark environment
6) incubated for 60min and removed plates and washed twice with PBS
7) replaced medium with PBS and recorded fluorescence on a multiwell
plate reader at ex485 & em520nm at time 0, 30 & 60min.
(I have also tried this assay by adding DCFH-DA followed by treating with H2O2)

I have observed a similar result in both ways, which was inconclusive. As per my knowledge, i should get higher values in treated cells
compared with non-treated cells. But i observed a reverse pattern for some unknown reason. Always untreated wells were showing higher values
than treated wells. Surprisingly the wells containing only PBS+DCFDA & without cells also showing similar fluorescence as untreated
wells(this shouldn't happen because this dye is non-fluorescent) .

I really appreciate your suggestions and recommendations in fixing the assay. I am mainly doubting my wavelength, are they correct wavelengths ? Do you mind sharing your protocol?

Thanks in advance,
Siva

miss montana on Mon Oct 19 03:09:54 2009 said: