I am trying to using Ni-NTA beads to do small-scale protein sample purifications (from cell-free reactions); however, majority of proteins still bind with sticky bead solutions after running western blot. May someone do similar process? and may provide suggestions on this issue?
Thanks!
Can you put details of how you are doing the binding, wash and elution from the Ni-NTA beads?
Stanley Ng Yes. Basically, I follow the instruction protocol. I equilibrium beads solution with equilibrium buffer (100 mM sodium phosphate, 600 mM sodium chloride, 0.05% Tween-20 and 30 mM imidazole, pH8) and collect beads. Next, I mix my sample with equilibrium buffer (1:1 volume ratio) and rotate for 30 min and collect beads. Then I wash beads twice with washing buffer (100 mM sodium phosphate, 600 mM sodium chloride, 0.05% Tween-20 and 50 mM imidazole, pH8). Finally, I add elution buffer into beads (100 mM sodium phosphate, 600 mM sodium chloride,250 mM imidazole, pH8) and rotate 15 min and collect elution samples.
Hope that is clear! Many thanks for the help!
Have you tried a higher molarity Imidazole elution, say 500mM?
And finish off with a EDTA strip, you can desalt the nickel out from your protein if needed.
This will establish if it's purifying via metal biding.
If not you're getting non-specific binding to the agarose beads.
You could try some his-magnetic beads instead, which tell you if it's the agarose.
Or blocking the beads with 3% BSA, like in an IP, to minimise nonspecific binding.
If the protein is binding strongly to the resin, you could try eluting the protein using a slightly higher imidazole concentration (~300 mM) at a lower pH (~7), as the affinity of the his tag for the metal will be decreased at a lower pH. Before trying this, you should ensure that the protein is stable at the lower pH first.
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