Dear all,
I may sound stupid enough.
I tried the PMS-DCPIP assay system as suggested, and I choose to observe the
absorption at 600nM. However, when I initialize the reaction, I actually see
the Abs slowly but steadily increasing rather than decreasing.
How the FAD interact with the enzyme is really unknown. No structure is
available. The only thing I know is that there is a FAD binding domain. I don't
know if it's dissociable or remain bound during the reaction
Another question might be how to properly dissolve lipid for assay. I currently
dissolve them using hexane and then add them to Tris-Chaps buffer (pH ~7, and
about 10mM Chaps). Is there a better way to dissolve them for assay.
Please advise.
michael nelson wrote:
> Dear all,
>
> Thank you for all your kind replies.
>
> Here is a little bit more about the enzyme and how I carry out the assay
at the first place.
>
> My enzyme is a lipid desaturase, originally from plant but overexpressed
in bacteria. FAD serves as a co-factor for this enzyme, in which FAD is reduced
to FADH2.
>
> My goal is set up an assay that would allows me to continuously monitor
the progress of the reaction. And I didn't want to use HPLC to analyze the
final product since that would take a lot time and we don't have an
instrument readily available to us. I wish FAD could be an alternative way since
FAD will have different Abs in reduced or oxidized forms.
>
> I set up assay is a regular lab setting (not anaerobic), add FAD,
substrate, ions and incubate. I finally add enzyme to initialize the reaction. I
expect to see some decrease of the Ab at 450 nM. But I didn't.
>
> I have several concerns, one is the autooxidisability of FAD, how fast
FADH2 would be reoxidized by O2 in the air or by the O2 dissolved in solution.
The second cocern is how fast the FAD reaction will go.
>
> Please advise.
>
> Thank you!
>
> Mike
>
>
