I'm going to express a protein in the membrane surface of a gram-negative bacteria and I wonder which should be the best protocol to obtain the membrane protein fraction for western blotting purposes. I understand there are in house protocols based in ultracentrifugation separation and also a few commercial kits available. Any recommendations?
This method that I am going to describe is tried and tested and works really well in my lab.
1. Once you have harvested your bacterial cells (over-expressing your membrane protein), wash the cells PBS to remove residual media. Re-suspend yor bacterial pellet (approx 4-5g wet cell weight) into ~ 30 mL lysis buffer*.
2. Lyse your cell suspension either via probe sonication (30 sec ON, 30 sec OFF, 12 cycles, keeping the suspension cool, i.e on ice) or French Press (3 passages).
3. Add appropriate levels of DNAse, RNAse, and protease inhibitors (I use the Roche inhibitor cocktail tablet). Centrifuge the cell slurry @ 27,000 xg at 4 deg for 45 mins.
4. Remove your supernatant and centrifuge at 120,000 xg at 4 deg for 90 mins (obviously using an ultracentrifugation (UC)). Remember to keep your samples cool (i.e. on ice at all times).
5. After UC, you should see a "dark brown/orange" residue at the bottom of your tube, this is your membrane fraction. Carefully remove the supernatant (cytoplasmic fraction) from the tube.
6. you now want to re-suspend your membrane fraction, this is the tricky part. If you mix this fraction up and down it will bubble due to the detergent effect of the lipids, this is not such as great idea, your activity will be affected. I find the best way is to completely remove all the supernatant using a P200 tip. Then add back to your membranes fresh buffer (what ever that might be, i.e most appropriate for your membrane(s) protein(s)). I usually add ~300 uL to the tube. Cover the top of the tube with some parafilm and leave on ice or at 4 deg overnight. The membrane residue will naturally "lift" into the solution and you will find it easier to re-suspend into solution via very gentle/mild pipetting action (remember not to introduce bubbles).
* remember, selecting the correct lysis buffer will dramatically affect the outcome of your experiment. This is something only you can answer.
This method that I am going to describe is tried and tested and works really well in my lab.
1. Once you have harvested your bacterial cells (over-expressing your membrane protein), wash the cells PBS to remove residual media. Re-suspend yor bacterial pellet (approx 4-5g wet cell weight) into ~ 30 mL lysis buffer*.
2. Lyse your cell suspension either via probe sonication (30 sec ON, 30 sec OFF, 12 cycles, keeping the suspension cool, i.e on ice) or French Press (3 passages).
3. Add appropriate levels of DNAse, RNAse, and protease inhibitors (I use the Roche inhibitor cocktail tablet). Centrifuge the cell slurry @ 27,000 xg at 4 deg for 45 mins.
4. Remove your supernatant and centrifuge at 120,000 xg at 4 deg for 90 mins (obviously using an ultracentrifugation (UC)). Remember to keep your samples cool (i.e. on ice at all times).
5. After UC, you should see a "dark brown/orange" residue at the bottom of your tube, this is your membrane fraction. Carefully remove the supernatant (cytoplasmic fraction) from the tube.
6. you now want to re-suspend your membrane fraction, this is the tricky part. If you mix this fraction up and down it will bubble due to the detergent effect of the lipids, this is not such as great idea, your activity will be affected. I find the best way is to completely remove all the supernatant using a P200 tip. Then add back to your membranes fresh buffer (what ever that might be, i.e most appropriate for your membrane(s) protein(s)). I usually add ~300 uL to the tube. Cover the top of the tube with some parafilm and leave on ice or at 4 deg overnight. The membrane residue will naturally "lift" into the solution and you will find it easier to re-suspend into solution via very gentle/mild pipetting action (remember not to introduce bubbles).
* remember, selecting the correct lysis buffer will dramatically affect the outcome of your experiment. This is something only you can answer.
To add to Luke Alderwick's thorough answer, I would recommend resuspending the membrane pellet in one of two ways, first having removed it from the centrifuge tube with e.g. a cell scraper.
a) Place in a Dounce homogenizer (https://en.wikipedia.org/wiki/Alexander_Dounce) with some lysis buffer and gently resuspend with several up-and-down strokes at a low velocity to avoid frothing/bubbling.
b) Draw the membrane and lysis buffer up into a syringe through a thin gauge needle (or first a larger gauge, followed by a thinner gauge) multiple times.
Both of these work well - the first is probably the easier of the two. Following resuspension, you can use the material directly for gels/blots, or you could solubilise the membrane with detergent and purify your protein if necessary, e.g. with an affinity tag if available.
The only problem with the protocols above is that if you follow them you will obtain an inner membrane fraction. This is why I ask for what kind of membrane Juliana has interest to purify and she answered she wants outer membrane. There is a protocol for the purification of outer membrane proteins published in: International Journal of Biological Macromolecules 39 (2006) 10–14 (Two-step purification of outer membrane proteins). In this protocol the differences in the detergent solubilization between outer and inner membrane protein is used in order to enrich outer membrane protein and then, and only then, to purify the outer membrane protein. This is a very important difference to be taken account because if you use the general protocol you will solubilize INNER membrane proteins and the outer membrane proteins (in which you have interest) will be lost in the pellet after detergent solubilization with the regular detergents. I myself tried this protocol and I have good results but in my case i work more frequently with membrane proteins from E coli inner membrane and Pichia membranes. Good luck!
In case you can't acces to the paper I mentioned above I am attaching it in this post. Good Luck
Thank you so much, Jose, Luke, Roger and Timothy for your thorough answers and gold tips! Thank you as well for the links and pdfs, you were of great help! The only protocol I got was not so well explained. I'll take everything in consideration while planning my next steps on the bench. Cheers!
Hello all, this a great thread with valuable information. I am following 95% of the protocol as suggested by Luke Alderwick. However, I am encountering certain problems. Here are my questions-
1. How do I show that my extraction is working by western blot? Could you suggest a good positive control for membrane protein so I can be sure there is no contamination of cytoplasmic protein? The reason I ask is because, I am trying to prove inner membrane localization of my protein of interest, but my protein sometimes goes to cytoplasm depending on growth conditions. I hypothesize this is a way to regulate its function (active in membrane, inactive in cytoplasm). So I need to absolutely make sure that my protein is the membrane at a tested condition.
2. What kind of buffer do you resuspend the pellet in? If so is it compatible for western blotting?
3. Does ultracentrifugation pellet just membrane fraction, or is there a possibility that large protein complexes (which I believe are also heavy) will pellet? Again, I ask because every cytoplasmic protein I tested (RNA polymerase alpha, adenylate cylcase, crp) hoping it would serve as a control, is strangely in the pellet (ie membrane fraction!) It's very confusing !!
Any recommendation will help.
Thank you.
Vidhya
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