[josenog]

Hi everyone!

I'm having a big trouble with a lipid-protein binding assay.

The protocol is quite simple:

1. Perform successive dilutions of a 5ug/uL lipid stock solution (I'm using two kinds of phosphatidic acid and phosphatidylcholine stocked at -80ºC on 50:50 MeOH:CHCl3 and use 2:1:0,8 MeOH:CHCl3:H2O for dilutions)

2. Apply 1uL of those solutions in Hybond-C nitrocellulose membrane (spots containing 5pg->5ug of lipid).

3. Let membrane dry O/N at RT protected from light. I already tried to dry it with N2 stream and with vacuum.

4. Block membrane with fatty acid free BSA (1h RT)

5. Incubate with protein of interest at concentrations 0,5-5ug/mL to test binding , (O/N 4ºC or 1-2h RT)

6. Wash membrane (5 times - 30minutes total)

7. Incubate with primary antibody (1h RT)

8. Wash membrane (5 times - 30minutes total)

9. Incubate with secondary antibody, conjugated with HRP (1h RT)

10. Wash membrane (5 times - 30minutes total)

11. ECL detection.


I already tried TBS, TBST (0,01-0,1%), PBS and PBST (0,01-0,1%) on wash, protein, antibody and block solutions.

I know the problem is not on antibodies/washes/ECL detection because regular Western blots with this protein always work.


I'm trying to repeat an assay that is already published and I know that this protein binds phosphatidic acid but not phosphatidylcholine.
What happens is that I never see spots...

Around the places where I spot the biggest amounts of lipid, sometimes it lookes like there is a darker area, stronger than normal noise (but it's not that evident).

Is it posible that lipid does not bind strong enough to the membrane and than just goes away on further steps?

Does anybody have experience with this kind of assay?

Thank you all for your answers!
Greetings