Hello Everyone, I´ve been working with microalgae and neutral lipid content; I´m trying to determine lipids in a 96-well plate using Nile Red dye but I´ve been facing some problems with the fluorescence results, anyone has experience on this??
I have been using Nile red for sometime now looking at neutral lipids in actinobacteria. You haven't said what problems you are facing, so it is hard to suggest the reasons why, however from my own experience, it may be due to the filter set you are using (Nile red has a blue shift in neutral enviroments), the nile red may not be pure; I have also noted a difference between staining with Nile red in aqueous and in an alcohol enviroment.
Hi !
I´m staining with Nile red (previously diluted with acetone) and DMSO (for membrane permeability), the problem is that I always get the same fluorescence results, they don´t increase or decrease with time, they are supposed to be increasing. (This is a protocol already reported).
I think the problem is the DMSO, which I recently realized is breaking the cells, and makes the fluorescence measurements go very high.
So my questions are,
Have you had experience with Nile red along with DMSO?
Does the breaking of the cells affects that much in the fluorescence result? Because, even though they break, the neutral lipids are released and stained.
What do you think?
Hi,
I have only used Nile red with ethanol, methanol and in aqueous. Speaking from experience, I was able to demonstrate that when the Nile red is dissolved in ethanol and applied to our bacteria, that the total signal was higher than with aqueous, but at a loss of definition of lipid deposits. I recommend initially dissolving the nile red in ethanol, then diluting it in water. Don't forget that nile red exhibits a blue shift in fluorescence depending on the enviroment it is in, so the released lipids may not exist in the same state as those inside of cells.
Finally, you could always try other molecules - BODIPPY for example.
Hi! thanks for your comments...
I´m now wondering about the concentrations that you use, since you dissolve Nile red in ethanol and then you dilute in water.
In the protocol I´m following, they dilute everything in DMSO, they also tried ethanol and obtained pretty good results, so I will try it as well.
But the protocol is very harsh on the cells (I think) because they suggest putting 5 ul of sample, 3 ul of nile red (50 ug/ml) and 292 ul of DMSO! I´m guessing it will be the same amount with ethanol, but I think it´s too much and I´m pretty certain that the cells will break again.
Would you suggest that instead of diluting everything in DMSO or ethanol I do it in aqueous?
PS. I haven´t read much about bodippy, I´ve only found articles where they only observe cells but they don´t quantify. I need to quantify in microplate... Any recommendations?
Thanks!
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