I need protein in Native condition.....For native I add DTT in by buffer for protein

Could it be that ionic strength needs to be adjusted? It could be that it is either too low, or too high. Both extremes favour precipitation. As a first approach, I would suggest you to try with a few different ionic strength values and see what happens.

Have you tried adding detergents to your protein before concentration?

The approach depends on what is causing the aggregation - most probably it is hydrophobic interaction. This could be countered by changing the pH some (not sure how pH sensitive your protein is) but a more likely approach would be to add a small amount of a biocompatiale detergent such as one of the Tweens - this should overcome interaction between hydrophobic areas of the protein.

I had this problem with a couple of variants of a protein I work with, I tried a range of PHs, different ionic strengths, different types of buffer and a range of salt concentrations. Finally, I tested a range of glycerol concentrations and adding 10% glycerol helped eliminate the problem.

Many proteins will aggregate when concentrated from solution. You have given very little information about your protein except that it forms oligomers or how you isolated it . Are these homo or hetero oligomers? Is the protein a membrane or cytosolic or even organelle linked protein. Have you included protease inhibitors?

Some things to consider:

Your salt concentration is pretty high - 2x physiological.

The functional pH of the protein gives some idea of the pH of the buffer you should be using. It should not be at the proteins pI as this is where the solubility is lowest and promotes aggregation and solubility problems.

Some chaotropic agents such as urea or guanidinium salts can help prevent aggregation but will need to be removed at a later stage to reactivate the protein (you will need to test the ability to reactivate the protein as some agents cause irreversible unfolding).

Some proteins use metal salts to aggregate and removal of these salts prior to concentration could help.

Addition of surfactants, glycerol or lipid derivatives could help for membrane associated proteins.

Removal of DNA or RNA could help if the protein binds to these nucleic acids.

I have experienced this before with membrane proteins. As a starting point, I agree you should check the isoelectric point (pI) of your protein and adjust the PH of your buffer accordingly. Add NaCl to stabilize your protein immediately after extraction; you can try different concentrations up to 400-500mM. I also have added 8M urea and up to 1M DTT alongside beta-mercaptoethanol at times to prevent disuphide bridge bond aggregation within cysteine rich regions of my protein. I have also tried 100% glycerol (v/v) and this prevented aggregation of my protein.

One important note, before adding any detergents or other chemicals make sure the membrane in the device you use for concentration is resistant to those chemicals. Some are more sensitive than others.

The Theoretical pI is 5.98 and average hydropathy value(-0.075) suggest, it hydrophilic in nature. When, I see its homo-oligomerization property using Galaxy gemini, I found that my protein forms 16 mer (homooligomer). That match with native PAGE analysis, as protein stuck in the well of 5% PAGE.....

I am agreeing with your suggestion that salt concentration is too high and also the pH...CAn you suggest me for buffer...as I purify protein at 4 dgree celcius...is I have to change the buffer from Tris to HEPES....as HEPES have some surface activity too and its pH is stable with temperature.

Need your valuable suggestions...Thank you all for suggestions

What usually works well is to pre-block the membrane of the concentrating device overnight with 5% detergent (tween/triton). Then wash the concentrator with your buffer and try to concentrate. Good luck!

I had similar problem, and I notice that adding 10% glycerol helped to decrease aggregation. You should also try other compounds that help to stabilize proteins

Different factors can lead to protein aggregation during concentration. The most evident is the concentration itself. Another one could be teh temperature, as at low temperature and depending of your buffer composition, you can observe aggregate. Sometimes, just by warming at 25 °C you can recovered the solubility of your protein.

The way to avoid precipitation could be to not concentrate too much your sample. You can also add low concentration of glycerol or detergents like NP40, Tween 20 or Triton X-100, which increase the solubility of proteins.

Sometime it is good to perform a buffer exchange before concentration in order to decrease salt and buffer concentrations.

If nothing works, you have to denature your protein with urea and to slowly renature it by successive dialysis.

Moreover if your protein of interest is a recombinant one, you can co-express it with chaperones (Gro ES, Gro EL,...) thanks to expression plasmid available in different firms.

Good luck.

As other sugestted the buffer that you are using contains quite high salt concentration, I would decrease to 20-50 mM and add 5-10% glycerol it may stabilize your protein. Does your protein contains His-Tag if so, removing his-tag will help with stability too.

Try adding a bit of EDTA and reducing agent to your buffer (mM not M!). Whether high salt or low salt is better is entirely variable from protein to protein, so you should try a salt/buffer matrix like what the crystallographers use if you have such a system available (usually a kit). We routinely use 10% glycerol to prevent aggregation during storage and sometimes use 0.2 to 0.4 M Arg or Pro. If you can't avoid aggregation maybe you can use it to your advantage to highly concentrate your protein, then dissolve the aggregates in urea or guanidine and do a dilution/dialysis refolding protocol to recover soluble active protein when you need it. Lots of literature on this (I know that this in itself could be a PhD project, but it's just a thought).

I had the same problem and I filled my diluted protein solution in a dialysis membrane and put the membrane in a dextrane polymer of appropriate kDa-size wetted with buffer solution. You have to work with three clamps to close and sequentially shorten the dialysis membrane. When the water is to approx. half of the volume drained out the membrane use the third clamp to shorten the tube for the next round of osmolarity-driven removal of liquid from the protein solution.

Try using glycerol (5%) during centrifugation step. Also try adding different aminoacids (Arginine).

I would suggest you to add glycerol to a final concentration of 5% before centrifugal filtration. HEPES would be the buffer of my choice. Will decide further after seeing the outcome.

Depending on what you want to do with your protein after concentration, you could try reduction + alkylation. With non-thiol reductant both steps can be carried out in one go,

Ultrafiltration may not be the best choice for your protein. An alternative is to dialyse against high molecular weight PEG. Use a high grade and high quality dialysis membrane. This method avoids the very high local protein concentrations that occur using ultrafiltration and can cause aggregation.

This has worked for several proteins: 50 mM arginine and 50 mM glutamate.

J Am Chem Soc. 2004 Jul 28;126(29):8933-9.

A simple method for improving protein solubility and long-term stability.

Golovanov AP, Hautbergue GM, Wilson SA, Lian LY.

The protein might be with some parttially folded segments, for this try also thermal shift assays with spyro green or Bis-ANS, to identify a better buffer condition which help the protein get more well folded.

Sometimes trehalose is usfull (10-15%).

I agree with Jörg Kleinschmidt, I have had the same problem with my protein and when I changed the pH > pI , the problem was resolved.

Amongst the other suggestions, I would suggest trying a non-denaturing zwiterionic detergent such as 10 mM CHAPS. I have enjoyed success many times with this in past purifications of difficult proteins that aggregate into oligomers. Keeping the concentration at or above the CMC (critical micelle concentration of approx. 6 mM) often allows aggregating proteins to be held apart even at high concentrations.

Dialysis

It may not help, but if you're using an Amicon ultra, try mixing the solution up and down every 3-5 min of concentration. During centrifugation, it often seems to get more concentrated at the membrane, so mixing it regularly may slow down oligomerization of your protein by keeping a more even concentration.

I think high salt concentration (see 'salting out') is the problem along with pI -pH relationship. Make the buffer at about 50 mM with low ionic strength and pH>pI or pH< pI over 1. Make sure that the pH should not change during concentration, if so adjust it accordingly.

I try diluting protein 3-4 times before conc step. Using 20 ml concentrators i concentrate it many steps. Each time taking out concentrate in separate tube and then proceeding with rest. Each time pool your concentrate separately Once you are done with all diluted protein start the concentrated elute. it takes time but it helps.

If your protein is fine at 10 C try the temp change.

An Amicon stirred cell concentrator will prevent a protein concentration gradient from forming during concentration which is likely causing the aggregation, see: http://www.millipore.com/userguides/tech1/99228

EMD Millipore offers a new device, Amicon® Pro Purification System, for multiple applications including general affinity purification, but it also transforms applications such as depletion of abundant proteins, protein enrichment, buffer exchange and clean-up of antibody labeling reactions. Overall, the System is extremely flexible and has a modular design so you can configure the perfect centrifugal purification device (i.e. Molecular weight cut off, MWCO) for your protein preparation. Search Amicon Pro http://www.millipore.com for additional information

For your application, simultaneous buffer exchange during concentration works gently so the proteins don't crash and can easily be recovered using a reverse spin.

Try to absorb solute with dry PEG 40,000 put onto dialysis bag with your staff.

Recommendation on buffer strongly depends upon protein nature.

You will succeed with adding 2-3 M urea if you proteins stands it.

Bhaskar Bhaskar ... Please check references below.. hope it will be helpful for you…

Regards…

http://wolfson.huji.ac.il/purification/PDF/Literature/Bondos2003.pdf

http://citeseerx.ist.psu.edu/viewdoc/download?doi=10.1.1.510.8414&rep=rep1&type=pdf