I am a PhD scholar working with HL-60 cells and I am trying to induce DNA damage to them and detect gamma H2AX proteins (15kDa) by western blotting using a well-documented toxin, Arsenic.
However, instead of seeing a band at the expected 15kDA region, I've detected nothing after repeating the experiment several times. However, my actin band appeared whenever I detected for H2AX and actin simultaneously but surprisingly, there were no band at all for H2AX...! Only once i got some light and merged bands at 15kDa but at that time no actin band appeared..!
I am very sure that the H2AX proteins are in the cells because I am using high molar concentration of arsenic that is sufficient to induce DNA damage.
where i am going wrong:
Running the gel at wrong voltage (100V) and time (2.5hrs)?
Using wrong PVDF membrane size (0.2uM )?
Transferring proteins at wrong current (200) or for wrong time (1h and 15min)
Using wrong dilution of primary antibody (1:500)?
Using wrong dilution of secondary antibody (1:10000)?
Here is my protocol:
1. Lyse the cells. centrifuge at top speed for 30mins at 4deg Celsius to remove cell debris.
2. Quantitate amount of proteins and load 20ul of protein sample onto a 13% gel. Run gel at 100V for 2.5hrs.
3. Transfer proteins to a 0.2uM PVDF membrane at1h and 15min.
4. Block membrane in freshly prepared 5% milk (in TTBS) at room temp for 1 hr.
5. Incubate membrane with anti-phospho-histone H2AX, diluted 1:500, overnight at 4deg Celsius.
6. Wash with TTBS (3X). Incubate with Rabbit HRP conjugated IgG (diluted 1:10000) for 1hr at room temp.
7. Wash with TTBS (3X) and detect with Immobilon Western Chemiluminescent HRP Substrate.
Please help me out with this problem of mine..
I shall be very thankful to you.
Edited by kokoakash, 13 August 2012 - 10:36 PM.