There are some steps that you might take to overcome this problem:

1) Add up to 20 mM of Imidazole in your sample and wash buffer. It can avoid weak interactions of His from other proteins

2) Make a linear gradient of imidazol in FPLC equipment. Try a "slow" (0,5 mL/min) 0-500 mM of Imidazole gradient

3) Adding 500 mM (or even 1M) of NaCl can avoid weak electrostatic interactions of other proteins with your protein or the column.

And keep in mind that:

1) The more the purity degree, the less yield you may achieve

2) This affinity purification is usually a first-step purification process. You can get higher purity using ion/size exclusion chromatography later (don't forget to dialyse your sample to remove Imidazole!).

Hope it helps! =)

As you say, it is a good way to start from 20mM Imidazole in your wash buffer to max of 50mM. Try doing them in parallel (20,30,40,50mM) from the batch input. Elute with same elution buffer and see the difference on the gel and pick the best!

Remember, you have to compromise some of your protein (yield) when you wish to have higher purity.

It is also a good practice to add imidazole to the sample and to the equilibration buffer to avoid the non-specific binding (5 to 15 mM).

Try including 0.5 M urea to disrupt non-specific interactions.

We use 20 mM imidazole+ 300 mM NaCl+ 5 mM B-ME throughout washing. Just before elution, wash with 10-15 ml of buffer containing 50 mM Imidazole + 300 mM Nacl, no B-ME. Taake elution with 200 mM imidazole + 150 mM Nacl. Usually 2-3rd fractions are richest.

indeed NaCl in your buffer is an excellent idea, I would even go up to 0.5M, add this + 20 mM imidazole in your sample before applying to the column and keep the same amount in your washing buffer. I would also go up with the elution concentration, 0.5M imidazole, some of my proteins only eluted at 300-400 mM imidazole.

You can use copper or cobalt instead of nickel.

Together with following above precautions optimize the elution in just 250 mM Imidazole or even 500 mM Imidazole if its enough.

Try non-Imidazol elution. His-tagged proteins can be eluted from the Ni-column using buffers with the pH below 7, without any Imidazole. Monomers usually will elute at pH around 5.7-6.0, while aggregates will elute mostly at pH around 4.3-4.8.

Have you tried to elute your protein doing a gradient of elution buffer? I understand that you are doing in-batch purification process. Using a FPLC you can perform a lineal gradient of elution buffer and separate the non-specific protein (hopefully eluting at lower imidazole concentration) from your protein of interest. Anyway putting 20mM of Imidazole in your wash buffer will considerably help you reducing non-specific proteins. In case you are doing in-bach purification process you can also try doing different washes with increasing imidazole concentration buffers in order to separate the elution of the non-specific proteins from the one of interest.

You have some nice suggestions above. Try to increase imidazol concentrations in wash buffers so you could determined the optimal concentrations to elute contaminants without loss of your protein. The use of continous gradient could help. The use of NaCl can give you some better results.

You can also add another purification step by either ion exchange or gel filtration in dependence of the contaminants. This could help eliminate imidazole and improve the final purity.

I would also maybe think about decreasing the bead bed volume. Too much resin per recombinant protein can lead to an increase in co-purifying contaminants.

Try 1% tween20 in binding and washing buffer along with imidazole as mentioned above. this might help in reducing the non-specific bands. Also you can reduce the binding time.

Another benefit to using the FPLC with a lineal gradient of elution buffer is that your chromatogram should give you some indication of the concentration of imidazole your protein can tolerate in the wash buffer. If a small scale run shows your protein eluting off around 100 mM imidazole then consider using a wash buffer with 20 - 50 mM imidazole to remove non-specific proteins prior to eluting you protein of interest.

There are some steps that you might take to overcome this problem:

1) Add up to 20 mM of Imidazole in your sample and wash buffer. It can avoid weak interactions of His from other proteins

2) Make a linear gradient of imidazol in FPLC equipment. Try a "slow" (0,5 mL/min) 0-500 mM of Imidazole gradient

3) Adding 500 mM (or even 1M) of NaCl can avoid weak electrostatic interactions of other proteins with your protein or the column.

And keep in mind that:

1) The more the purity degree, the less yield you may achieve

2) This affinity purification is usually a first-step purification process. You can get higher purity using ion/size exclusion chromatography later (don't forget to dialyse your sample to remove Imidazole!).

Hope it helps! =)

Maybe in FPLC step, you should collect small amount of eluent in each fraction size. Only 0.5 ml or 0.1ml if you need it. And you can use step gradient elution for protein purification. i think you should not use concentration of imidazole higher than 300 mM if you said it reduce your protein yield. you should optimize the imidazole concentration between 200-300 mM. If all not succeed, keep thinking about the second purification. Good luck ^_^

Prewash thoroughly with NaCl and weak imidazole; may be you can also try 0,01% triton or NP-40in the first wash. Later you can exclude detergent

better do it as an assay, so u can decide the optimum concentration..also run the fractions of washings too in the same gel. Must check both stained gel and the blot.

use 10% glycerol in your buffers.

increase the salt concentration to 0.75 M in your buffer

try using cobalt resin

cheers

As Fabricio said, you should keep in mind that affinity purification, and in particular Ni-NTA, is generally the first step of a purification process due to its low specificity. In my opinion, the most important advice for getting higher purity are 1) adding 20 to 40 mM imidazole in your sample (don't forget to check until how much imidazole concentration the protein remains bound) and 2) working with at least 500 mM NaCl in all your buffers including the sample buffer.

If you express the protein in fusion with a solubilizing partner, there is a way to significantly improve purity using only Ni-NTA. You first purify the fusion protein. This fusion protein should contain a protease cleavage site between the target protein and the partner. Cleave the fusion protein by adding the protease (you may change the buffer before). To separate the 6-His partner from your target protein, load again the sample on the same Ni-NTA column (don't forget to add 20 mM imidazole in the sample). The main advantage of this strategy is that both contaminants and 6-His partner will bind again the column whereas the target protein will not. I found this strategy very efficient and inexpensive (in particular if you express and purify the protease by yourself). On-column cleavage also works, but purity was not so good.

To minimize the contaminats in Ni-NTA purification, There are two standard ways.

1. As everyone above was suggesting, include some imidazole (may be upto 50 mM, if your protein does not elute at this) in loading and/or wash buffers to prevent lose binders.

2. Decrease the salt concentration, to prevent the chaperone interactions and co-elutions, since the common contaminants are the chaperones GroEL (60 kD), DnaK (70 kD) and DnaJ (40 kD).

Good point about chaperones...

May be addition of ATP 5mM and Mg 1mM in all buffers might help to get rid of Chaperones..

El siguiente articulo te podría ayudar a desarrollar una estrategia:

Bartlow et al., 2011. Identification of native Escherichia coli BL21 (DE3) proteins that bind to immobilized metal affinity chromatography under high imidazole conditions and use of 2D-DIGE to evaluate contamination pools with respect to recombinant protein expression level. Protein expression and purification. 78: 216-224. doi:10.1016/j.pep.2011.04.021

Reducing the amount of the Ni-NTA resins might also help: since there is a sort of competition for Ni2+ binding between the recombinant protein, cellular proteins with naturally encoded histidines and free imidazole, less affinity matrix will provide more specifically bound proteins, i.e. his-tagged ones. Of course, there is the issue to optimize the amount of resins depending on the yield of the produced protein. Elution of the desired protein at different conc of imidazole might also infer that the population of the protein is not entirely homogenous, so I'd suggest checking for the activity of eluted fractions. I had a protein that would partially elute during the wash step (40 mM imidazole) but the fraction had poor activity and I discarded it assuming incorrect folding. The more is not necessarily the better.

You can reduce the amount of NI-NTA matrix..sometimes if you take more amount of NI-NTA matrix there is a possibility of impurities in eluted protein..so try it take around 1 or 1.5 ml of NI-NTA matrix..may be this can solve your problem..

gud luck.

hi

If your system Ni-nTA for the QIA Gene company you could Use a better answer you will get.Performance is Bach system more of the Column. Moreover, if one sequence of His-Tag the concentrationOf imidazole is 30 mM enough if the two His-Tag sequences in your protein, you must 75 mM imidazole Used in wash buffer.

Addition of imidazole up to 20_30mM consentration to wash buffer can avoid binding of non specific proteins to Ni. In addition reduction in amount of resin can be usefull too. Because if there is resin_Ni more than needing of his tagged protein , non specific proteins can bind them.

From your qusetion it seem like you are not adding Imidazole in your sample during binding. Try to add 10-15 mM of Imidazole in your sample before loading into Ni-NTA resin.

As many researchers suggested about reducing Ni NTA matrix amount , i would like to ask if we we are using pre-packed HIS trap FF 5ml column from GE then how can we optimize the elution? 

To reduce the non-specific binding first determine at which imidazole concentration your protein elutes. Then take the maximal imidazole concentration where your protein does not elute and extensively wash the NTA with this.

This usually removes all the weak binding bands.

It is rare that a protein can be purified to homogeneity in a single step. Try ion exchange column after Ni-NTA.

There are some tips I would recommend to decrease elution of non-specific protein with your target. Based on my experience, you have to include imidazole in the both binding and washing buffer and in your starting sample in a concentration that do not elute your target protein (I usually use 20-50 mM imidazole in both starting sample, binding and wash buffer). Second is to Increase NaCl concentration, use at least 0.5 M in binding buffer to prevent nonspecific binding to IMAC