Timir, several people have made very good general suggestions, but you might get better help if you provided some additional information. Most people are probably assuming that you are expressing your protein in E. coli. Is that correct? This point is relevant because your extraction method may be contributing to the problem. You might be able to adjust your extraction conditions to favor release of your expressed protein preferentially over the contaminants. Also, you should consider the possibility that your contaminant proteins are actually binding not to the Ni-NTA column, but instead are binding adventitiously to your expressed protein. Have you tried running an extract of cells that that are not expressing your protein over the column? If you do that and don't see the contaminant bands, your problem will not be solved by modifications aimed at decreasing the binding of the contaminants to the Ni resin. Again, adjusting your extraction conditions could help minimize adventitious binding to your expressed protein. But you could also disrupt such interactions with chaotropes such as Hoffmeister salts (sodium iodide, sodium bromide) or other large monovalent anions such as thiocyanate, trinitrophenolate, or trichloroacetate. This is old school protein purification technique that is very effective, but the knowledge is sadly becoming lost because it's generally so easy now to purify tagged, overexpressed proteins.

His-tag clones are always impure when purified with Ni-NTA. Normally I used to purify it with a second round of chromatography. Since in your case it is mainly high mol wt impurity, try running size exlcusion column after running it through Ni-NTA.

In first instance you can try a imidazole gradient, that might give you some fractions with higher purity. However, I also would recomend an extra round of purification. You may try Q or S sepharose. The easiest way is under conditions where your protein is not bound, so you can take the flowthrough and bind directly to the Ni-NTA.

Another "trick" can be using pre-packed niquel columns, like HisTrap, which have higher binding capacity, so you can try higher imidazol concentration during the binding.

In any case, from my experience, I would advice other tag, like GST or Strep-tags. Those tags are much more specific than histidines.

In addition to the already posted comments: try to elute with a pH shift instead of imidazole, but make sure that you do not use a pH range, which denatures your protein of interest.

Hi Timir,

two quick thoughts :

1- is your protein well expressed ? I mean is it 1mg/L culture or 50 ? It makes a big difference with His-tag purification as it is said above specificity is reather low.

Use Talon to increase specificity or Machery Nagel Ni-TED resine (care cause it is silica based and flow is quite different). Moreover, elution under much more fractions implying you dilute more your protein when eluting.

2-use strep-tag very specific my prefered choice.

good luck

you could try TE pH 7.4 as lysis buffer, Try to keep step gradient after washing.

Can you lower your pH to around 7.4? If your protein is leaking at 40mM, then the binding is not as tight. As others have suggested, Imidazole gradient might help. Other alternatives are Co2+ containing matrix e.g TALON.

If you have access to an FPLC, thats the easiest way to sort out your impurities. Many proteins in bacteria can bind to the Nickel column due to histidine exposure in their tertiary structure. On an FPLC, you will be able to run an imidazole linear gradient and collect the specific fractions where your protein elutes out. A subsequent PAGE/ western blot should tell you the exact concentration of imidazole at which your protein elutes out. Sometimes using a higher imidazole concentration allows the elution of more proteins than you are interested in.

Alternatively, if you dont have access to an FPLC, I would suggest that you elute your protein by increasing the imidazole concentration in small steps like 50mM, 60mM so on and so forth instead of a big jump to 200mM. You can then concentrate your protein from the different fractions if necessary

I would recommend size exclusion chromatography post NiNTA purification to get rid of the comtaminants, sine NiNTA purification does not most of the time yield pure protein.

As others have suggested, I would recommend trying Talon resin as this has better specificity towards his tag. Another 'trick' I can think of is to reduce the volume of the resin, because the more resin you have, the more nonspecific binding will occur.

Two points: wash the column with many volumes of 0.5 M and 1 M NaCl or KCl in phosphate, no imidazole, then elute. Second: it is not generally appreciated that PMSF is not a significant protease inhibitor in alkaline buffers; it degrades from base catalyzed hydrolysis faster than it inhibits even chymotrypsin. DFP is much more effective and not nearly as toxic, is spite of its reputation.

As others suggested above:

- try to start the purification with an ion-exchange chromatography step that would work under similar condition as Ni-NTA chromatography does,

- combine the imidasole elution with pH shift (for instance, I do binding and washing at pH 7.8 and 10 mM imidazole and elution at pH 7.2 and 250 mM imidazole).

What is the nature of the "contaminating" bands? Are these bands on silver-stained gel (or on a Coomassie-stained gel) or on the Western blot (or on both)?

One more thing; if contaminating bands are at higher MW than your requested protein, try to finish your purification protocol with a size-exclusion chromatography step.

I think you can increase the imidazole conc in wash buffer. Yes you will loose some of your protein but the result will be better.

2nd thing use MW cut off column to carry a size-exclusion chromatography step.

I hope this two steps will give you high quality protein.

First thing is to try use lower temp for protein lysis in SDS Page to 95-98d

I would suggest to use ion exchange or size exclusion chromatography. In addition you can cleave the tag by Thrombin or TEV and then bind the protein again on the Ni beads. Collect the flow through and you will remove all the contamination bands (That will still be bound to the Ni beads)

I would suggest to optimize the expression condition if you haven't done. The more protein expressed the less impurities from column, Also as Lee suggested, reducing the volume of resin will help lower nonspecific binding.

Has anyone tried NiCo21(DE3) from NEB ?

Hi Timir,

I went through most of the suggestion, and am sure that it will help you optimise the purification protocol.

I have worked with most of the His-Tagged resins, and i would say it does not matter what resin you use. It does not change thing that much.

I would rather focus on your expression sysytem, and try to optimise it there. if you could give the details of the Bacterial expression sysytem and conditions used to express you desired protein, i would be in better position to comment.

Also you cannot have the pure protein in a single purification, and would likely need a 2-step purification system. Affinity followed by IEC or HIC. RPC is also good choice for 2 step if activity and structure are not your focus.

You could also check the empirical property protein using the amino acid sequence (eg amino acid composition %age, chagre, solubility etc). This along with the final use of your protein, will be very important to decide your second purification step.

hope to hear from you

Use 30-50 mM imidazole+ 500 mM NaCl in binding buffer. Adjust pH to 7.5 for best binding with Ni-NTA resin. You dont need to wash the resin with different conc of imidazole containing buffer. Just wash 5-10 column of binding buffer then elute with 200 mM imidazole containing buffer.

Try washing the column with the same washing buffer with little salts also like 200 to 300 mM NaCl. Also increasing the number of washings till A280~0 would also work. Could you use Tris buffer????????

Timir, several people have made very good general suggestions, but you might get better help if you provided some additional information. Most people are probably assuming that you are expressing your protein in E. coli. Is that correct? This point is relevant because your extraction method may be contributing to the problem. You might be able to adjust your extraction conditions to favor release of your expressed protein preferentially over the contaminants. Also, you should consider the possibility that your contaminant proteins are actually binding not to the Ni-NTA column, but instead are binding adventitiously to your expressed protein. Have you tried running an extract of cells that that are not expressing your protein over the column? If you do that and don't see the contaminant bands, your problem will not be solved by modifications aimed at decreasing the binding of the contaminants to the Ni resin. Again, adjusting your extraction conditions could help minimize adventitious binding to your expressed protein. But you could also disrupt such interactions with chaotropes such as Hoffmeister salts (sodium iodide, sodium bromide) or other large monovalent anions such as thiocyanate, trinitrophenolate, or trichloroacetate. This is old school protein purification technique that is very effective, but the knowledge is sadly becoming lost because it's generally so easy now to purify tagged, overexpressed proteins.

Daniel made a very significant point here : the possibility that some E. coli proteins interact with your protein of interest (although in our experience no more than one or two proteins were revealed to interact with the overexpressed one). Mass spect allowed their identification. They were GTP binding proteins and adding GTP to the sample before Ni-NTA loading released the interactions and in turn contamination.

i agree with both Daniel and Thierry!!

However i would say that Protein purification in not that easy task as it looks. Having the details of your expression system and protein would etch you some good troubleshoot tips. But i would also recommend Daniel's advice, as it will tell you if the impurities are from your leaky expression or something that interacts with your protein.

I have Mass spec analysed many proteins which actually bind to column because of leaky expression that bind to Ni-resin, like SlyD at 23/28kDa and few other proteins at 99.67; 43.28; 66.91; 51.56;74.29 kDa. They have high affinity for Ni-NTA, and most of then can be eluted at abt 60mM imidazole but then you risk of eluting your protein as well.

Streptag is realy worth considering. Ok you have to clone again. I suggest you start to share with the "Gateway community". Many Invitrogen inspired plasmids have been customized.Homologous recombination renders cloning procedures less problematic sometimes.

Instead of using a gradient for washing and eluting at a constant imidazole concentration (200 mM) you should apply a gradient for the elution. Also, 40 mM imidazole for washing appears to be relatively high, in particular, since your protein starts to come off at 40 mM. By the way, did this fraction with your protein contain the same amount of impurities as the 200 mM elution fraction? Anyway, I suggest washing with 20 mM imidazole and using a gradient of 20 -200 mM imidazole for elution. You may obtain fractions enriched with your protein without the described HMW impurities.

I have tried several different His-tag resins, the Talon resin works better in that there is lower non-specific binding, but they way you achieve that is by using less resin than is needed to bind all your His-tagged protein. In this way, your His-tagged protein will compete off all the other weakly binding proteins (batch mode binding is the way to go with Talon, columns did not work well for me). But you can only wash Talon with ~10 mM imidazole since there is a weaker interaction with the His-tag (at least in my case). An initial gradient elution is good to find where your protein comes off, then you can step to it on a big prep.

There's been a TON of great chromatography suggestions for you. Excellent suggestions. One thing people haven't really commented on is perhaps what you're seeing on your SDS-PAGE. I have a couple of points for you.

You say your protein is a 50kda dimer? Are you expressing a fusion of the monomer, or does it have an affinity for it self? Maybe provide a little more info about your particular protein and the down-stream analysis you intend to use :) . That'll get you more suggestions.

SDS-PAGE points to consider.

1. NEVER take an SDS-PAGE for face-value! People do this all to often. If you see one 50kDa band, and nothing else, great, you've got protein >95% pure. If you see multiple bands, you may have your protein really pure and free of other proteins, but what you are seeing is a combination of dimers, trimers, tetramers, etc of your dimer (that would give you a 100-200kDa weights, etc.) People think typically give too much credit to their PAGE sample buffer (SB) for denaturing their samples, and assume all is working "textbook"--not the case. You CAN have dimers, etc in your SDS-PAGE is your protein is not reduced enough in your SB (is it old). Add excess B-mercaptoethanol. SDS-PAGE gels make a lot of ROS (reactive oxygen species) from the electrolysis of your running buffer (which is why you see bubbles in the tank when you start your SDS-PAGE). These oxidize met and cys residues. Try adding an antioxidant to the inner chamber of your running buffer. You can buy little bottles of running buffer antioxidant from Life (Invitrogen).

2. Find an epitope in your protein, and verify whether of these higher-weight band contain the same epitope as your protein. You do this by western blot. Yes, it's a bit of work to optimize, but it's a good way to to give you a second dimension of analysis. Make sure you run your Ecoli lysate on the western blot to make sure your primary antibody isn't immunoreactive with other proteins in the lysate.

3. Life (Invitrogen) sells SDS-PAGE stains that have a specificity to staining proteins with His-tags. Consider this before blotting (much cheaper and faster). If any of the heavy-bands stain really well, there's a good chance they have the His-Tag you've put on them, and thus are dimers/trimers of your expressed protein.

Suggestions for His-Tag purification

Not sure what your protein is for, but nobody (that I read) suggested a denatured lysis/or binding to the Ni-NTA resin. This obviously ins't the first choice, but if purity is an issue, consider it. Denaturing in urea or guanidinium will eliminate these potential interactions, because proteins more-often-than-not depend on their tertiary conformation for binding interactions (specific or not). Once you've washed everything away, you can refold on the Ni-NTA resin. Or, you can elute in a denatured state and refold by either rapid dilution (very specific protocols and tricky!), dialysis, or on a gel-filtration/size-exclusion column. If you don't have an FPLC, you can do the former with gravity-flow as well.

I purify prions, so I KNOW a "tricky" protein. I adopt the attitude "Don't give your protein too much credit in getting you down!" A protein is a protein is a protein. When in doubt, get aggressive, then work backwards! ;)

Good luck,

It's hard to judge without seeing an image of the gel or knowing the nature of your protein, but one possibility is that what you are obtaining is a mixture of different truncates of your protein due to digestion by proteases. Are you using protease inhibitors during extraction?

Hello Everyone,

I am having a similar kind of problem purifying my protein of interest.I am expressing my protein of interest with a fusion peptide and a 6X his tag,the total size being around 22 KDa.After purifying through the Ni NTA there is a contaminating band around 50 Kda on SDS-PAGE.I thought it could be a dimer so I incubated it with BMe and did SDS-PAGE once again but this 50 KDa band was still present.So I used sephadex G 100 to get rid of this contaminating protein but after gel filtration when I did SDS-PAGE and silver staning I could see very very faint bands of my protein with 50 Kda band also there.I dont understand why this happened and how could I RESOVE THIS PROBLEM?

use an His tag antiboy to perform a western blot. You will see if the 50 KDa is tagged and related to your protein. Wired things may happen. We experimented homodomers which were resistant to SDS-BMe reducing conditions. Also in special conditions covalent Lys-lys or Tyr-Tyr cross-linking may occure between strongly interacting proteins like elastin, insect resilin, plant extensin, collagen. Mass spec may help.

Thanks sir

Hi!

There are always serious things to be considered for getting rid of the protein contaminants. But here I would like to suggest some very simple things. Follow them if possible and see if that improves your purification process.

1. extract in higher volume of buffer. 1 to 5 @ w/v concentration, i.e., 1 gram pellet should be dissolved to at least 5 ml of buffer. if using lysozyme mediated treatment then incubate for 20 min on ice and then centrifuge to collect the supernatant or folow mechanical means to get the protein in soluble phase. use nuclease to reduce the viscosity of the solution.

2. add imidazole to the protein solution to be purified @ 10mM final concentration. this effectively reduces contaminants from binding to the column. elute at half volume of the starting material. in this way two washes (one with 20 mM and the other 40 mM) are good enough to reduce or remove the contaminating proteins

We routinely are facing problems of this kind in the lab. The main problem seems to be that the amount of the protein of interest is low in the extract. Consequently, a significant number of sites on the Ni column can be occupied by less specific binding. You can reduce the amount of resin to be used but below 300 µL of slury things become diffcult to handle. You can perform a second elution after removing imidazole of the first purification. You can use NiTED (Machery Nagel) which is more specific but give you less yield and elute in a larger volume (too bad if you need a concentrated protein fraction). People are usualy polishing with gel exclusion chromatography but again you dilute the sample. The lesson : nothing's perfect but you already know that.

Have you resolved this problem yet?  I would say an additional 1 or 2 chromatography steps  are usually required.  One suggestion that I did not see in any of the comments (my apologies if I missed it) is the use of a reducing agent in the binding/wash/elution buffers.  It is common to include a reducing agent such as TCEP or DTT to reduce intermolecular disulfide bonds.  It is possible that your contaminating proteins are forming disulfide bonds with your His-taged protein causing them to be co-purified.

Hi all,

Same in my case, I assume if you try two step purification would help

Such as,

Remove the various other proteins using Q-Sepahrarose Ion exchange chromatography followed by His tag purification on NiNTA/Talon

This (2 step) worked for me when my protein was secreted into the media

I'm currently working on various combinations of purification from cells (Sf9)

Have fun.....All the best

Raju

hi.. Timir.. i think, there are some interacting proteins which elutes with your portein.

i suggest you to reduce the sonication time and also increase the salt concentration 500 mM in your buffers.

 How I can avoid to binding histidine rich bacterial protein to NINTA column during purification of recombinant protein in E.coli? THANKS

@Karolina: It is a common problem, especially when your protein of interest is expressing lower (lets say a eukaryotic protein expressed in bacteria). I would recommend you to equilibrate and bind your sample with 20-30mM Imidazole; wash with same buffer containing 400 mM KCl, 2.5 mM MgCl2 and 1 mM ATP to ger rid of E.coli chaperones. I know it wont remove the 100% of your contaminants but it greatly reduce! some people(me too) use wash buffer cotaining 0.5 to 1Molar NaCl! Please make sure which is suitable for your protein of interest (you have to compromise a part but not everything).

Hi,

I don't know if somebody already suggested or not but have you tried ATP wash?. This can really help you getting rid of many impurities. You can use 1-5 mM in wash buffer (make fresh just before use). 

Good luck 

How many cysteines?  Are you using a reducing agent?  TCEP is very effective and will not reduce Ni, although compatibility with phosphate might be am issue.  I didn't see a reducing agent listed so maybe this protein has none.  Buffer choice can affect aggregation,  make sure your picked your buffer and pH based on the isoelectric point of your protein of interest.  Many comprehensive suggestions here, good luck.