Hi Ramon,

something worth noting is that if you are exchanging with 4L overnight, you can get a much more potent dialysis of the imidazole by using the same 4L, but in 4x 1L steps.

ie - say you have 50ml protein at 500mM Imidazole to start with. Dialysis against 4L overnight will result in a final concentration of 6.25mM.

2x dialysis against 2L will result in an intermediate imidazole concentration of 12.5mM, and a final concentration of 312.5uM.

4x dialysis against 1L => 25mM / 1.25mM / 62.5uM / 3.1uM - i.e., a 2000-fold greater dilution of the imidazole with the same volume of dialysis buffer. This is also a lot more effective than diluting the protein sample (160,000-fold dilution would be necessary to get the same decrease in Imidazole concentration : this may cause some problems with handling 8000L of solution in subsequent processing steps!)....

Good luck! Robin

Dialysis will only save you one day (not hands on, you can do something else in between). Dilution can be an option if you are sure that imidazole is not going to interfere withfurther experiments you pa to do with your protein.

It depends on your following experiments. If your protein uses His for interactions or catalysis then Imidazole is a problem. To get rid of Imidazole I either dialyse into assay buffer 3x100 fold dilution (total of 10^6 dilution) or simply run a size exclusion/buffer change column that I have equlibrated while doing my metal affinity purification (you can overload the column quite a bit when the size difference is that large)

If you are extremely pressed for time you can always check if the presence of imidazole is an issue and do all your assays in imidazole buffers.

Seems relative as far as my experience is concerned. 3 of the proteins i worked with had no effect on the type of method i used for removing imdz (diluting or using dialysis bag). However, other proteins behaved well when dialysed over night and gave consistent readings in successive assays or experiments. So i always check the compatibility of protein towards the two methods.

We always do at least one more purification step after IMAC. Most often an ion exchange step. We often use 150 mM NaCl in the IMAC buffers (at least for elution - sometimes more salt is required for binding/washing) and just do a dilution and pH adjustment before the ion exchange.

We always carry out a thorough dialysis after any purification, anything else labeled as 'timesaving' is just laziness. As stated above you can get on with other things while its in dialysis and regular changes of the dialysis buffer should be done. Its a gentle technique (rather than centrifugal concentration followed by dilution) and you can change buffer types as you go to avoid any possible problems of precipitants forming (something you may well observe in the through wash when washing your column even before applying your protein. I stongly recommend you do this if you are going to follow up with crystallisation trials of your protein, or carry out sensitive enzyme assays. Even the smallest, low concentration of contaminants can really mess you up.

Does saving a day really help?

Depends a lot on the situation. We develop processes for small scale protein production and dialysis is out of the toolbox for us. If buffer exchange is required we do it by chromatography, but if it can be replaced with dilution we prefer that. For final formulation we always do buffer exchange. In some cases I prefer mixed mode (e.g. Capto adhere) after IMAC and one of the reasons is that you can often load the IMAC eluate as is.

While it is similar to buffer dilution...and there is a benefit to a dilute protein preparation...one approach we've used which works quite well is the use of Spin Concentrators with a 100 kDa (or an exclusion size which keeps your protein from going through) exclusion membrane. You can dilute your sample buffer, spin to a desired volume, re-add more buffer, spin, etc. until you have decreased your Imidazole concentration to the level you desire in the final solution. This also allows you the ability to easily add other buffer constituents.

Another means for doing this, my preferred option, is spin-desalting columns. In this case, you use a Sephadex G-50 equilibrated in the final buffer of choice. You add your protein to the top of the gel after the void volume has been 'spun out' at 1000 rpm for 1 minute (typical table top swinging bucket centrifuge...used for cell sedimentation, etc.). You then spin your sample through the column at 1000 rpm for 1 minute. This desalting process fully exchanges the initial buffer with the bed-volume buffer in one quick step and, what is nice, without any protein dilution step. Protein concentration remains the same as does the input volume...provided all the void volume of the support was spun out.

Enjoy the fun.

One other factor to consider is protein stability. If your protein is not very stable in high salt and imidazole, then you want to change the conditions as quickly as possible. Then dilution of your protein solution (if necessary) and loading onto a second column would allow you to change buffer conditions rapidly. I find that a single Ni column step is not usually sufficient to get acceptably pure protein.

I prefer using a desalting column or buffer exchange with concentrators to save time. It's not just about laziness. You also reduce the amount of time your protein is in a high concentration of imidazole, which can be destabilizing. Also, you often can elute from a nickel affinity column with much lower than 500 mM imidazole, often 100-200 mM.