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Troubleshooting protein purification cation IEX - Cation IEX (Sep/23/2008 )

Dear All,

This is with reference to the purification of our recombinant protein sample expressed in E.coli as inclusion bodies. After Solubilization refolding we perform the cation exchange chromatography of our protein sample using SP sepharose fast flow resin. Attached herewith find the SDS PAGE and IEF results of the collected fractions.

In addition to our protein of interest we are also getting high molecular weigh contaminants, which we cannot get rid of in IEX. Can anyone please guide me on a technique to get rid of these bands as even after gel filtration of samples few high mol wt contaminant bands are not separated from main proteins and sample gets diluted too.

In cation IEX procedure is
Column Sp Sepharose Fast flow packed in fineline 35 column packed bed volume 100 ml
Equilibration 10 C.V. protein loading 3-4 C.V. [approx 1mg/ml protein conc], washing to remove unbound materials 2 C.V. step elution 0-35% gradient – 1 C.V., 35 –80% gradient – 10 C.V. and 60 –100% gradient 1 C.V.
Protein elutes at 40-50% gradient.
Protein details: Our protein is stable at acidic pH and has a pI of 5.8 –6.3 and buffer is Na Acetate buffer pH 4.5 and elution buffer is starting buffer containing 0.4 M NaCl.

We get only one peak on AKTA but on running SDS page we get so many bands even IEF shows 1-2 bands at the most.

How can we modify the method or what can be done to get rid of extra high mol wt bands.

Any help will be deeply appreciated.


A couple of things:
Was your pI calculated using denatured protein? If so, the real pI of the folded protein might be significantly different.
Try purifying by anion exchange at high pH.
You can test both anion and cation media to see the optimal conditions for binding and elution. Take loose media and equilibrate in a range of pHs. Add your sample batch-wise and mix for 10 minutes by gentle inverting. Spin the beads out in a microfuge (<1000 xg) then remove the supernatant. Run beads and S/N on SDS PAGE (don't forget to boil the beads!), and you'll find the best pH range and media for binding and eluting (it will be the lowest couple of pHs where all of the protein binds-that is, nothing in the S/N). Next, mix some of your sample with more beads equilibrated at the optimal pH; spin the beads and remove the S/N. Split this into a number of tubes and add some buffer with different amounts of salt. Run again on a gel, and you have the optimal [NaCl] for elution.