I have carried out a His-tag assisted affinity purification of a protein (43kDa) and now I am trying to buffer exchange and concentrate down my protein sample using a Vivaspin 20ml concentrator (30K MWCO) to concentrate down my protein. However it takes too long for the buffer to pass through the filter (1h-1h30min). Does anyone know whether this signifies that something is wrong with the protein sample?
Thanks in advance for your time!
P.S. I know I should have used a concentrator with lower MWCO but I wanted to get rid off a 27kDa protein that co-purifies with my protein.
We produce large amounts of protein for structural studies and they usually need to be concentrated to anywhere between 4 to 30 mg/ml. Concentration runs of one to two days are not uncommon for us, so I wouldn't worry about spending two hours concentrating your protein. We set our centrifuges to 4oC and spin them at 4000 rpm.
As has already been mentioned, resuspending the protein every 15-30 mins is helpful as this stops it from forming a gradient. The protein at the bottom of the concentrator will eventually become much more concentrated than the portion at the top and it can easily crash out of solution.
Also, to reduce the probability of your protein sticking to the concentrator membrane, you should choose concentrators with nylon membranes or at least PES (polyethersulfone) membranes, but stay away from any membrane that has regenerated cellulose in it.
This maybe due to membrane fouling where your protein particles settle on and block the membrane pores. Visually note the difference between unused and just used membrane. Probably a solution is just to half fill the 20 ml for concentration run.
1 to 2h does not seem so long to me. It depends how much volume you concentrate and what the concentration of your protein is. The more protein you have, the slower it will be. Check if your protein is concentrating as it should or if you lose it. Check the absorbance of the flow through also. Check also if you see any precipitation. Is this a soluble or a membrane protein? Do you have any detergent, glycerol or other more viscous components in your buffer? This will also contribute. Good luck!
Thanks to everyone for taking the time to answer my question.
After further investigation I found out that protein aggregates were building up on top of the membrane and this is probably what was causing the delay of flow through. I found this out by using my pippette to pippet up and down close to the membrane and then spinning down the supernatant in an eppendorf tube. This resulted in precipitate appearing at the bottom of the eppendorf.
Usually for us also it takes more time like yours. You can centrifuge it for short term like 10 mins then take it out and mix it nicely with pipette then put it back for centrifuge. repeat this step after 10 mins. it helps you to concentrate your protein fast then usual. still it will take bit long time so have patience and let it be concentrate at around 3000 rpm.
We produce large amounts of protein for structural studies and they usually need to be concentrated to anywhere between 4 to 30 mg/ml. Concentration runs of one to two days are not uncommon for us, so I wouldn't worry about spending two hours concentrating your protein. We set our centrifuges to 4oC and spin them at 4000 rpm.
As has already been mentioned, resuspending the protein every 15-30 mins is helpful as this stops it from forming a gradient. The protein at the bottom of the concentrator will eventually become much more concentrated than the portion at the top and it can easily crash out of solution.
Also, to reduce the probability of your protein sticking to the concentrator membrane, you should choose concentrators with nylon membranes or at least PES (polyethersulfone) membranes, but stay away from any membrane that has regenerated cellulose in it.
As mentioned before aggregation is mostly the reason for slow concentration, when it is starting at low concentrations.
Centrifugation, and 0,22 µm filtration and repeted mixing can help.
But one other point: If you want to get rid of a smaler protein (27 kDa using a 30MWCO) you may see that its not working with such a small difference in size: by technical reasons theres a variation in the pore size of the filter membrane ( at least 10%) - in my experience proteins often also tend to dimerizise and therefore "negative" filtration mostly does not work as expected.
I would speculate that since the manufacturer's recommend using a MWCO of 1/3 of the MW of your protein (as native state proteins can be more compact), you might actually be pulling some of your protein into the pores which would explain the blockage, also I'm not sure how well this system lends itself for actual protein seperation. I'd recommend looking into a size-exclusion chromography column for that.
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