I am trying to purify a protein whose pI is around 9 and I am using cation exchanger (Mono-S) with buffer of pH 7. But, my protein always comes out in unbound fraction. I tried using Mono Q as well thinking theoretical pI may be far off than actual pI but, in vain.
Also, 260 absorbance of my protein always comes as double that of 280 despite it being non DNA binder. I tried doing Dnase treatment by adding Dnase in lysis buffer but, still no change.
Can anyone please suggest what should I do to
1. Purify it by ion exchange
2. Reduce 260 absorbance.
It may be that your protein is badly aggregated. This would tend to explain the lack of ion exchange column binding as well as the greater absorbance at shorter wavelength (caused by light scattering). You can test this by using dynamic light scattering or size exclusion chromatography.
Hey Adam,
I did Size Exclusion Chromatography (SEC) of my samples and its not coming in void volume (If it would have been aggregated, it should have come in void volume, right ??) but, its not even coming at its expected size i.e around 58da rather its coming out around 90kDa which is also a bit strange !
Totally confused as to what is going wrong with my experiments !!
Hi Chitra,
I think Adam hit the nail and you should try DLS. What buffer did you use for both IEX and SEC? Did you maybe use (more) salt in the SEC? Some proteins tend to aggregate in low salt conditions and dissociate at a certain salt concentration, maybe your buffer/conditions of the SEC induced partial "re-dissociation" of your protein-aggregate. Your protein - if aggregated - will elute later during SEC. What is the protein source? Is it possible that there is another protein present in your solution? Maybe a binding protein like a chaperone? What do you see in a SDS-gel (Band at 58 kDa or higher weight?)? Nevertheless, I think it is worth to perform DLS (maybe under different conditions)!
Some proteins strongly absorb at 260 nm (but I don´t think this should lead to a doubling of the absorption at this wavelenght)
Best Regards,
Aleks
A mass estimate of 90 kDa for a 58 kDa protein by SEC is not a cause for concern because an elongated shape of the protein can cause it to appear larger by SEC. It is encouraging that the protein did not elute in the void volume, since this is consistent with it not being aggregated. This still leaves the puzzle of why it did not bind ion exchange columns. Something I should have asked before is, what is the ionic strength of the buffer the protein sample was in when you tried to bind it to IEX columns? If it was too high, this could keep the protein from binding. To lower it, either dilute the protein with a low ionic strength buffer, or dialyze it against a low ionic strength buffer.
Thanks for your reply Adam and Aleksandar !!
1. Ionic strength of buffers that i am using are very low. Roughly around 50mM Tris and 10mM NaCl in low salt buffer and 1M NaCl in high salt buffer
2. Its a fungal protein that has been expressed in E.coli Rosetta DE3 cells.
3. There are no contaminating bands with my protein.
What does very low exactly mean? Does it lead to a conductivity of 0.1, 0.5, 1 or >10 mS/cm??? Did you estimate it or is it a guess based on your buffer content? Because 50 mM TRIS can be very critical, especially when considering how it was prepared .... did you use TRIS-base or TRIS/HCl to preapre your buffer? If you have used TRIS-HCl then your buffer will exhibit high conductivity and the salt will extremly compete with your protein during IEX binding! I always use a maximum of 10-20 mM TRIS (no additional salt, if the protein can handle this) and check the conductivity to be at ca. 1 mS/cm!
Not knowing about the fungal protein amino acid sequence, is it soluble at pH 5-6? The surface exposed charge groups are what you have to deal with and in case the effective pI is not 9 then try to go to an extreme with your mono S with pH 5-6 acetate with no added NaCl to see if there is firstly binding and then if there's no precipitation at the head of the column assuming your protein is correctly folded. The lower pH will give stronger binding in the presence of higher conductivity as well as slow oxidation of cysteines. If you get bind/elute behavior the you can go to higher pH and lower conductivity. This can be done quickly at a tiny scale batch-wise using SP ImpRes even though it's a different strong cation exchange resin. Does it require reducing conditions? Is there an activity assay? You're at the edge of Tris HCl buffering capacity at pH 7 even at 50 mM.
Go lower in pH on the cation exchange as Grant suggested and no NaCl in the binding buffer. Tris is not an optimal buffer for cation exchange as it can interact with the ligand. In the column you pH may be far from 7.
You may have better luck with a mixed mode resin such as Capto adhere or MMC
"3. There are no contaminating bands with my protein."
Then why do you want more purification steps?
Aleksandar : Conductivity of my buffer is around 2.5mS/cm and I use Tris-HCl.
Grant and Bo : I can try that . Thanks for your suggestion !!
Bo : My third statement meant "Major band is of my protein" (around 90%) and rest one / two other bands
So I think you have now some options to try! 2.5 mS/cm can already be critically high (depending on the protein), so what I would try is - as already suggested - go further down with your pH (if your protein is able to handle this low pH) and use a buffer with a maximum conductivity of 1 mS/cm (use another buffer/or keep the pH and the TRIS buffer but use TRIS-base and lower concentrations of TRIS)..... For example I worked with a protein which bound the colum at 0.9 mS/cm but failed at 1.2 mS/cm (depending on its charged surface residue distribution) ..... Good luck with your work!
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