I am treating RAW 264.7 macrophages with some plant extracts dissolved in ethanol and measure if they decrease ROS/NO production. I have been having some trouble measuring ROS.
A condensed protocol is as follows;
1) Seed 500K cells, in 500ul volume per well in a 24 well plate 37C and 5% CO2. Let them adhere for 3-4 hours.
2) Add 2.5 ul of H2DCFDA 5mg/mL per well
3) Incubate for 30 minute 37C and 5% CO2.
4) Aspirate media, wash cells twice with warm PBS.
5) Add 500ul DMEM media.
6) Add 1ul of plant extract with [50mg/mL]
7) Incubate for 1 hour at 37C and 5% CO2.
8) Add 10ul of LPS [100ug/mL]
9) Incubate over night (18-20 hours) 37C and 5% CO2.
10) Read fluorescence in that same plate at 485nm excitation and 520nm emission in a plate reader.
Somehow the numbers are all over the place. Can anyone suggest improvements? How do you measure ROS in your lab? Should I change the incubation time?
Any feedback is appreciated and I would be more than happy to clarify/expand on the protocol details,
I followed the following protocol and it works.
https://www.abcam.com/dcfda--h2dcfda-cellular-ros-assay-kit-ab113851.html
My lab uses the Griess reagent to measure nitrite production. I'm not sure if that's better, but it is very easy and consistent for RAW 264.7 macrophages, and can be done in a 96-well format.
I think you may wish to also be exact in your LPS incubation time; pick 18 or 20 hours, this should remove some of the variability.
First suggestion for you is try to use flow cytometry instead of plate reader. Since you loaded your cells with DCFDA at the beginning, it may be OK for 6 to 12 hours treatment but 20 hours may be too much. You can try reading your plates earlier than 20 hours. I also worked extensively with plant compounds and in my own experience ROS generation is maximum at 4-9 hours (depending on compound). Good luck
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