I have purified a membrane protein via affinity purification using detergent OG. I was trying to dialyze the protein with ST (NaCl-150 mM and Tris-20 mM) buffer(pH 8). Unfortunately, during dialysis, the protein gets precipitated inside the dialysis bag.
The predicted pI of the protein is five, so I also tried the pH of the buffer adjusted below and above the PI. But no luck; it is still being precipitated. Any suggestions and answers will be helpful
Did you have any detergent in the dialysis bath? Although OG is not too easy to remove by dialysis, its concentration may have dropped below the minimum necessary to keep the protein in solution.
You need something to favor the solubilization of the membrane protein, as Adam B Shapiro comments, some OG might come out of the membrane during dialysis. The ST buffer that you mention will not work for a membrane protein. What is the initial concentration of OG?
If OG means octyl glycoside, it has a micelle size of about 8000 Da and a high CMC. it is therefore relatively easy to dialyze away. Indeed it can be used for reconstitution of membrane proteins into liposomes because of this property. Loss of (some of) the detergent is almost certainly the reason for your aggregation.
It seems as though you have no detergent in your dialysis buffer. As OG is pretty expensive I can see why you might not want to add it. What you may wish to consider is adding a non-dialyzable detergent to your sample prior to dialysis. You may have to screen a few detergents in order to find something that can replace OG and maintain protein function.
@Nick That makes sense.You are right about not using the OG,it is quite expensive.
If I need to have detergent in the dialysis buffer is it okay to use any detergent which has a lower MW ?
A way to minimize the amount of OG required to perform the dialysis with the detergent in the dialysis bath is to keep the volume of the dialysis bath to the smallest practical volume. Instead of preparing, say, one large 1-liter bath, you might instead use two sequential 200-ml baths, requiring a total of only 400 ml. It takes more time but saves on detergent.
There are certainly less expensive detergents than OG, but it can become complicated to replace one detergent with another in an existing sample. For example, you might decide to use an inexpensive polyoxyethylene-type detergent such as Triton X-100, Tween-20, or Brij-35 to replace OG during the dialysis step. However, micelles of these detergents are too large to dialyze, so they must be added to the sample. This results in mixed micelles of OG and the other detergent, which have properties different from either detergent. In the end, you don't know the final detergent composition of the sample.
You can also consider eliminating the dialysis step entirely if it isn't essential. If you need to change the buffer composition, you can use a centrifugal concentrator. You first concentrate the sample, then dilute it with the desired buffer. Repeat once more to get a more thorough replacement. A caveat is that the OG micelles may not pass readily through the ultrafiltration membrane, so the OG also concentrates along with the protein.
There are some detergents that make small micelles that can readily pass through ultrafiltration membranes. An example is CHAPS. If you use this detergent instead of OG, you can use the concentration/dilution technique to perform buffer exchange, without concentrating the detergent. Just remember to keep the CHAPS in the buffer that you add back when re-diluting.
If you start with one detergent and want to change to another, the best time to do it is while the protein is bound to a column. You can simply wash the column with a buffer containing the second detergent.
This paper has an example of this approach.
https://www.jbc.org/article/S0021-9258(17)41923-5/pdf
When you purify solubilised membrane proteins, the detergent used needs to be present at the cmc in all purification steps. This ensures that the amount of detergent in the mixed detergent/lipid/protein micelles does not change. A decrease would lead to precipitation, an increase to removal of anular lipids. Note that the cmc depends on buffer composition and needs to be determined for each buffer you use.
Dhanashree Lokesh , you need detergent to be at a sufficient concentration at all times to prevent aggregation. Detergent added to the dialysis buffer is only of value if it can enter the sample and replace the OG that is exiting. In your case it is not yet clear which detergent will actually prevent aggregation.
The molecular weight of the putative replacement for OG is not that relevant- most detergents are less than 1500 m.w. The CMC (the concentration at which micelles form) and the size of the micelle (varies by detergent type) are the critical parameters.
Any detergent with a small micelle size relative to the 1OK dialysis membrane will pass through. This is happening with OG hence you are losing it.
Often the detergent micelle is much larger than 10K and if the CMC is low and esp. if it is below the working concentration of your detergent, it is almost impossible to remove the detergent by dialysis as it exists only as large micelles. Of the commonly used detergents, CHAPS easily passes through dialysis membranes and Triton is almost totally excluded.
Thus in order to use Triton you would have to add it to your OG-containing protein sample prior to dialysis or work into your purification procedure as discussed by Adam B Shapiro .
Good Day :)
I recommend going away from detergents altogether. Membrane proteins are tricky, but not impossible to get solubilized.
The traditional detergent based approach has the problem that you need to constantly add detergents to the samples, no matter what stage of the purification step.
Also, the membrane protein gets removed from its native lipid environment. And even the best detergents are almost guaranteed to interfere with the membrane protein to some degree.
The more modern approach are nanodiscs, especially the ones based on copolymers like Amphipolds or SMA. They leave the surrounding lipid environment intact AND you do not constantly have to add them.
This should be of use for you:
https://cube-biotech.com/what-are-nanodiscs-synthetic-polymers
J. Stolz By replacing them with the copolymers like DIBMA, SMA, or AASTY.
See this page about SMA for example. Especially figure 4:
https://cube-biotech.com/smalp-features
The copolymer forms the stabilizing nanodisc and solubilized the membrane protein at the same time. No need for a detergent micelle.
If one insists on doing the solubilization still with detergents, MSP nanodiscs are also a possibility. See this page:
https://cube-biotech.com/what-are-nanodiscs-the-basics#mspnanodiscs
Also, figure 4.
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