I have a maltose binding fusion protein being purified with the MBPTrap column (GE). I get very little protein (less than 1 mg from a 400 mL E. coli BL21 (DE3) culture) during the elution step with 10 mM maltose buffer, however there is another large peak eluting during the wash step with NaOH. Running this fraction on a SDS-PAGE gel I can see a lot of unwanted, tightly bounded proteins as expected but also a lot of my target fusion protein.
Is there a limit on the amount of maltose that can be used for the elution step in this column? The protocol says a isocratic step at 10 mM is enough for elution but perhaps a gradient up to, lets say 500 mM might help with the recovery but I don't know if this good for the column before I try it.
I've seen that some people use up to 50mM of Maltose (could anyone confirm?). I haven´t tested it yet and I wouldnt recommend it. Whenever I got problems with my protein concentration what I try first is test protein production (optimize the best media, incubation time and temperature) temperature plays a big role on protein solubility (Low temperature seems to help to some proteins. Then I test sonication time and amplitude and/or pH values of both: lysis and elution buffer (I was having some problems as well, I prepared new Lysis and Elution buffers, measured pH with a better pH meter and then I got the expected protein concentration).
I would think that the concentration of maltose is not your problem. In my experience it takes very little maltose to elute from the dextrin-sepharose. On occation I have run a gradient elution and the fusion protein always comes right in the beginning of the gradient.
I assume you are using MBP fusion because you had solubility problems? MBP is a wonderful tool capable of solving many solubility problems, but sometimes it is just making your aggregated protein float. I have seen soluble MBP-fusions that should have been around 60 kDa run in the void volume on a Superdex 200 column!
I think that what you are seeing is aggregates that have precipitated on the column and are getting dissolved with the NaOH. If you have the possibility try running an analytical gel filtration to see if your protein is the expected size. If you see aggregates try adding 1-2M Urea and/or 10-20% glycerol to your material going into the dextrin column.
I use a gradient of 20-200mM maltose. Usually the protein elutes in the beginning of the gradient, but you can try if your protein needs more concentration of maltose. Also, as suggested look if your protein is forming aggregates.
1-2M Urea usually would not denature the protein. I do not mean this for cleaning of the column but for the actual purification. If possible you should try to do an analytical gel filtration to determine if aggregates are present or not. Usually MBP fusion is used because expression without MBP yields only insoluble protein. The risk is that your target protein is still not correctly folded, but the MBP will keep it in solution anyway.
I use amylose resin to purify MBP-fusion proteins.
On PAGE, I analyse the crude extract (column load), a sample of the unbound fraction, a sample of the "late unbound" peak (to check that the column was not overloaded) and samples from the maltose elution peak.
I often see the target protein in the unbound fraction, but te amount is trivial in comparison with what is eluted by 10mM maltose.
While the NaOH eluted material might well be aggregates, can you guesstimate how much of your protein is present in this fraction, versus how much was in the crude extract/unpound peak?
It might simply be that your protein is not well expressed in E.coli.
As you get material eluted with 10mM maltose, I cannot see how increasing its concentration will help, unless somehow the MBP has been mutated to a higher affinity version??
If indeed you have aggregation, as we have ocassionally seen, adding or increasing dithiothreitol (up to 10mM - provided your protein is not affected!), mercaptoethanol or TCEP might help to reduce this.
- Badges
- Science method