Hi,
I ve been stuck by the in vitro yeast Atg8 lipidation assays for a long time. I followed the basic protocol presented by Ohsumi's group [ Ichimura, Y. et al, (2004). In vivo and in vitro reconstitution of Atg8 conjugation essential for autophagy. Journal of Biological Chemistry, 279(39), 40584-40592. ].
The question is when I mixed Atg7 (2 uM), Atg8G116 (5 uM), ATP (1 mM), MgCl2 (1 mM), and DTT (1 mM), and incubated the reaction at 30゜C for 1 hr, the bands of Atg8 on gel was rather smear and became very similar to the lipidated form of Atg8 (i.e., lower than the original unmodified Atg8 bands). However, if ATP is omitted or Atg7 is absent (whatever Atg3 was present or not), the bands was as normal as the unmodified Atg8.
I suspect that it is the Atg8-AMP intermediates as the paper reported [ Nath, S. et al, (2014). Lipidation of the LC3/GABARAP family of autophagy proteins relies on a membrane-curvature-sensing domain in Atg3. Nature cell biology, 16(5), 415. ]. But anyway, this phenomenon really troubled me so much, since the smeared Atg8 bands hindered me to detect the 'REAL' lipidated Atg8. It caused a false appearance that I could not distinguish whether the lipidation reaction is normal.
By the way, the PAGE details are as follow:
- Separating gel: 12% (29:1 Acr:Bis) with 6M Urea;
- Stacking gel: 4% without Urea;
- Runnning buffer: Tris-Tricine electrophoresis buffer;
- Loading buffer: 5x-SDS loading buffer (with DTT);
- Boiling: 95 ゜C for 10 min.
So could anyone give some instructions or comments for the assays? Many thanks!
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