Good Day,

I hope this can help:

These are our protocols and buffer compositions when purifying a his tagged protein with Ni-NTA MagBeads. I unfortunalety cannot tell you a possible why this precipitation happens to you. But maybe switching to our native purification protocol solves this isse:

https://cube-biotech.com/purification-of-his-tagged-proteins-under-native/denaturing-conditions-using-purecube-his-affinity-magbeads

What colour your precipitate is? Have you tried to put the rest of the supernatant in a SDS PAGE? It confirms that your protein is precipitating?

Hey Choi,

As Jorgaq already pointed out, the first thing to do is to identify if the precipitation is indeed your protein. Since the solution is empty according to the Bradford assay, I would assume that your protein is in the precipitate - but running a gel can't harm.

There are a number of things you can try out.

Firstly, did you equilibrate the beads before adding them? If the beads are left in their storage slurry, you are introducing aqueous ethanol solution into your condition, which could trigger osmotic shock. Always make sure that you equilibrate your beads with water, then with your buffer to ensure that you do not cause osmotic shock.

Secondly, You can try to increase the salt concentration in your buffer. You are at pretty low salt concentration, so salting in may help the solubility of your protein. Beware that going over a certain threshold of salt can have the opposite effect and your protein will precipitate, so maybe try something in the 100mM-300mM range.

You can also try to increase the viscosity of your buffer by adding Glycerol. In my personal experience, this has helped my proteins stay in solution more easily. Go for something in the range of 2.5%-10% and see how it behaves.

Of course, you can also look at the construct design. If your construct is missing parts of the C- or N-terminal sequence, they may contribute to the solubility of your protein. Cofactors that may have been identified in previous studies should also be considered, and you may need to introduce the cofactors during your protocol to ensure the stability of your protocol.

Lastly, you can look into solubility tags. I do not have any experience with re-folding solubility tagged proteins, but I don't see how this would be impossible.

Thank you all! I almost gave up about getting answers... That is why I find your answers today. :)

Jorgaq and Michael,

1. Precipitates are white, and I confirmed that precipitates are my target protein by specific antibodies.

2. Beads are equilibrated with refolding buffer which I use. Precipitates are formed about 20 minutes after incubation begins.

3. I should try to add more salt, thanks.

4. The buffer contains 20% of glycerol, so I guess this concentration is enough!

I have doubts about the construct of my plasmid, but I cannot find ant information to resolve it. I consider that preparing a new plasmid. Currently, I am trying to purify this protein in the denaturing condition. Based on that the precipitates are formed about 20 minutes after incubation, I assumed that the major cause is the agitation. Furthermore, in fact, the pI of this protein is predicted to 7.6, close to the pH of my refolding buffer, I think that this might cause precipitation when agitation began.

Thank you all, again.

Hey Choi,

Glad I could try to give you some pointers. I don't know what kind of yields you are getting from lysis prior to your affinity purification, but you may want to go back to the drawing board and design a few buffer conditions that you can screen through in order to arrive at something suitable.

If you must keep your current construct, or if you want to try to optimize your purification before you re-design the construct, you could try making some small-scale purification (e.g. if you use 5mL of your bead slurry for one whole batch, you could try 10 different conditions at 500uL each).

How did you come up with the buffer in the first place? Has there been prior success in purifying your specific protein using this buffer system?

There are a variety of factors that can affect protein solubility, but the four major factors I would consider in your situation are:

1. Salt concentration - As mentioned above, both "salting in" and "salting out" can have a beneficial effect. Try to play around by making some with high salt and maybe one low salt condition compared to your current buffer.

2. Isoelectric point - Now with your pI at 7.6, your current pH is fine as long as you do not plan to perform any ion exchange chromatography purification. You will need to be at least off by 1 point on the scale compared to your pI. Here, you can consider which environment your protein would be found natively? Is this an enzyme usually found in an acidic or basic environment?

3. Viscosity - I should have noticed the fact that your buffer contains 20% Glycerol, so I apologize for not catching that earlier. I don't see why this specific concentration would cause you any problems, but you can try to play with something slightly lower, maybe in the 5-10% range and see how that affects your solubility.

4. Cofactors - This can very much make or break your buffer. You are working with a Kinase, which means that the obvious additive here is Magnesium, which you have in your buffer. But the kinase may have binding partners that it requires to be happy, or other small molecules you aren't currently supplying. If you are working with an active kinase and you have some other way to produce reasonably pure proteins using an antibody-based approach, one way to screen how "happy" the protein is in its current buffer is by performing phosphorylation activity assays. You only need tiny quantities of protein, and you can test a great number of conditions. There are even commercial buffer screens that you can get for this very purpose. The higher the activity, the better the buffer.

I hope you find a way to overcome this obstacle.

Choi Wonseok, i agree pretty much with all that Michael says. It seems weird that your protein aggregates just after adding Ni beads. If you think that agitation causes a problem, try to agitate your protein solution before adding beads (even though 12 rpm is not a lot).

Otherwise, do you really need 10 mM of Beta ME in your buffer? Are you sure that Ni resin can hold all of that? Furthermore, you can try to add a little bit of a detergent to see if that makes a difference or not for the solubility of your protein.