I work with a protein which shows cooperativity on kinetic assays, and others of its class (although with low similarity) are dimers. However, it's not described in literature for this one that I work. There is no crystal ou NMR structure solved.
To determine this we already tried DLS (Dynamic Light Sccatering), but the predicted dymer doesn't induces a big size change compared to the monomer, so it's an inconclusive test. We tried gel electrophoresis (Native PAGE), but the protein buffer contains Triton, which forms micelles, making also inconclusive. Triton also interferes on SAX (Small Angle X-Ray diffraction) and size based chomatography... Is there any other technic that I can use where Triton micelles doesn't interfere??
Analytical ultracentrifugation (AU) by sedimentation velocity (SV) analysis is one of the classic methods for determining the oligomerization state of a protein. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2373601/
If I get you right, you are talking about 'dimer' not 'dymer'. If that's the case, gel filtration chromatography can help to resolve whether your protein is dimer or not. if Triton is interfering, you can easily remove it by using either spin columns or by dialysis. if the dimerization is due to the disulfide bonds, denaturing SDS PAGE with and without BME/DTT would give some idea.
You can try chemical crosslinking, using a low crosslinker concentration, followed by SDS-PAGE. Please see the attached paper for an example.
Easiest way is to just do a native (nondenaturing) gel electrophoresis (with controls, of course), and see if there is a band corresponding to the dimer. Computationally (if structure is known), analyze the protein surface to see if there are complementary patches. If you want to know if it functions dimerically in vivo, use nanoscopy (ultramicroscopy) methods, of which several are now available.
Few more ideas. From your explanation I concluded that you have Triton micelles involving the dimeric protein. If, I'm right, then remove Triton from buffer (by dialysis over night for instance) and then run SDS-PAGE and native gels. IF your dimeric protein is within the micelle you have high probability of getting monomer as soon as triton is removed.
On the other hand there are several gel filtration columns already prepared to work with Triton-buffers. From my experience, this approach is relatively cheap and quick.
Rosa is right in that the Triton needs to be removed, at least down to the cmc. This can be done easily with hydrophobic gels like Biobeads SM2 (doi:10.1016/0003-2697(73)90436-3, doi:10.1590/S0100-879X2002000700001). The resulting solution can then be investigated further:
- Analytical ultracentrifugation is the gold standard. If you do not have access to an analytical instrument, you can use the procedure of Martin & Ames (http://www.jbc.org/content/236/5/1372.full.pdf+html?sid=cc0a124d-c88f-431b-90b3-3570a52e8ca2). This can even be performed in a desktop centrifuge (doi:10.1016/0003-2697(89)90297-2). However, a real analytical instrument offers many advantages (doi:10.1110/ps.0207702).
- Native electrophoresis, perhaps best the blue variety (doi:10.1006/abio.1994.111).
- Gel filtration. Note that if your protein was solubilised with Triton, the running buffer needs to contain Triton at the cmc to prevent protein aggregation. SEC columns need to be equillibrated with detergent for several days before you can add sample. The cmc depends on the buffer composition and temperature, it is easy to determine (doi:10.1016/0003-2697(84)90026-5).
- Laser, neutron and X-ray scattering should work once the excess Triton has been removed.
You need to determine with what links the dimer is formed. If using a hydrophobic interaction then it is not a dimer. This is an associate. If using disulfide bonds, then it is a dimer. Destroy the hydrophobic interaction can be solutions of urea, ethyl alcohol. Disulfide or sulfide bonds under these conditions will remain.
Even non-covalently bound couples of membrane proteins are dissociated at 2% SDS-60*C. If protein subunits are covalently bound vis S-S linkage, you can examine beta-ME or DTT treatment. I agree with proposed above action of urea, formamide, DMSO, etc.
P.S. ..and then the Laemli' PAG-electrophoresis is the best way to solve the problem.
Quick way I have resolved dimerization detection is to run the the same sample on SDS-Page gels in consecutive lanes under +/- denaturing condition. And B-Me or DTT containing loading dye to 1st sample and No B-Me or DTT to the last 2 samples. Run a control if available and MW Marker, then +, -, -,. You should see a shift from the monomer ( in the + lane), a distinctive split profile from monomer to the dimer( the 1st (-) lane) and the dimer in the last lane. Sorry no example to show.
Hi Pedro,
As already mentioned above by Rosa and Engelbert Gel filtration/SEC with or without the removal of Triton would be the simplest way to go.
I would also suggest you another neat way: I assume that you have your gene with a tag for purification. In case that you have it with two different tags (or even one with a tag and one without one) you could in principle perform immuno-precipitation using the first tag and after you run the gel, detect your protein with an antibody against the second tag: if you get only one species with the IPed tag then your protein is most likely a monomer. However, if you find the protein that has the second tag (that was not used to IP the protein), it will indicate that the protein is in a dimeric form.
In case that the tags adds a substantial weight that affects the mobility of the protein on the gel, then even a single tag should be enough if the two forms of the expressed protein can be detected on an SDS-PAGE or by western blot.
Analytical ultracentrifugation (AU) by sedimentation velocity (SV) analysis is one of the classic methods for determining the oligomerization state of a protein. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC2373601/
Though not widely known, the existence of dimers or other oligomeric forms is readily assessed via spin labeling and electron paramagnetic resonance spectroscopy. We've employed it with great success in assessing the existence of multiple forms of oligomeric aBeta in solution, work to be published in the spring of 2019. The method is amenable to biologically relevant media and relies wholly on changes in basic physical properties. FYI, our results do not conform to those from gel separation methods, nor does DLS work for the molecules we've examined.
Size exclusion chromatography without TritonX in the buffer is probably your best bet.
Do you have access to SEC-MALS instrumentation with UV and RI monitoring? You can do a conjugation analysis and obtain the mass fractions of detergent and protein within an eluted peak, and then use the MALS signal to determine the mass of your protein and thus obtain the oligomeric state. The method is often used to characterize detergent-solubilized membrane proteins, have a look at this: https://www.rug.nl/research/portal/files/6726538/2008MethodsSlotboom.pdf
You can try crosslinking. Crosslink your protein with an available crosslinked and then run an SDS PAGE of the sample. Now this may or may not work. If you have the means try Size exclusion Chromatography. This works the best.
Analytical centrifugation can be the option however Size Exclusion chromatography can be used as Triton didn't interfered when i purified my
protein having Triton in the buffer.
Diffusion NMR (DOSY) can be used and should be effective for proteins at 1 mg/ml concentration. Triton will give strong background signals but there should still be areas where the protein can be analyzed to give a diffusion coefficient. You do need to run some individual protein standards to get a calibration.
Standard 5 mm NMR tubes are probably not the best. It would be good to use tube inserts. Traditionally, the tube insert contains D2O but it is handy to put the sample in the tube insert and the D2O in the tube.
I published a basic protocol a long time ago ( Analytical biochemistry 331 (2), 395-397, 2004 ) but you will need some modifications ( Polymer Chemistry 8 (44), 6700-6708 is useful). DOSY is an established method at USP-chemistry (http://labrmn.iqm.unicamp.br/). I'd be happy to consult with NMR colleagues to adapt any local protocols for protein analysis.
Regarding chemical crosslinking (mentioned by Ditsa Sarkar ).
This method appears simple and straight forward. It involves some simple chemistry involving a bifunctional small molecule that can react with free amines (Lys sidechains), carboxylates (Glu/Asp sidechains) or sulfhydryls (Cys sidechains).
Like all chemical reactions, the relative concentration of the protein and crosslinking agent, as well as time and temperature, are important factors in the reaction. What is the optimal conditions? Too dilute, short time, low temperature and you get no crosslinked dimer product. Too concentrated, long time, higher temperatures and you can see non-specific dimers. These aspects are rarely mentioned or discussed in papers.
While the aim might be to crosslink two amines in the two parts of the dimer, in reality, you most often crosslink two amines in the same monomer unit. The result is that when you do electrophoresis, you get an envelope of protein products:
monomer, monomer plus one crosslinker, monomer plus two crosslinkers etc.
So, the higher the crosslinking agent, the broader your bands will be in electrophoresis. This is a limiting factor in achieving a 100% yield of crosslinked dimer.
Few people do chemical crosslinking with proper controls. My personal experience was that I had to do about 30 test conditions to optimize the reaction conditions. We also used known monomeric and dimeric proteins as controls that I just didn't see in the literature back in 2000 (I suspect it hasn't changed). It was impossible to find conditions where the control monomer gave 95% dimer.
For these reasons, chemical crosslinking is one of my least favorite methods for checking monomer/dimer questions. It is a fast experiment that takes months to optimize. See European Journal of Biochemistry (2001) 268 (23), 6229-6237
Hi.....i think I can determine by electrophrasis of this protein in presence of standerd protein
Hi. I think that by studying different methods for extracting proteins, extraction and sequencing can be done correctly. Different methods in the world of berry protein extraction are used
In my opinion, analytical centrifugation and/or size exclusion chromatography may be useful
For a globular protein, a dimer should be 2^(1/3) ~1.25 of the size of the monomer.. That size change should be observable, however may be challenging in the presence of micelle interference.
Here is a range of typical protein DLS sizes versus molecular weight
https://www.materials-talks.com/blog/2016/01/21/zetasizer-sensitivity-for-protein-dls/
The simpliest method is running a SDS-PAGE under non-reductive conditions. Make the normal process with a dye without beta-mercapthoethanol and do not heat the samples. This will be enough to see if the protein has a dimeric conformation. I have done it, and got good results.
I agree with Pamela Pena and have used this method. You should run SDS-PAGE under non-reductive conditions.and you should use sample dye without beta-mercapthoethanol and do not heat the samples. This will be enough to see if the protein has a dimeric conformation.
Yeah, you may proceed with non reducing SDS PAGE as experts have suggested above. Additionally, you prefer to sequence your protein. Furthermore, if you know the gene sequence, which coded your protein (if you have been heterologously expressing it), then an analysis using a software, which predicts the protein production could help you identify the exact molecular weight ( I am not sure if many researchers would recommend this)
As I wrote earlier:
1) Take the definite mass of the purified protein under study;
2) Calculate the number of its micromoles;
3) Proceed analysis of the N-terminal amino acid residues (there are several suitable methods) in micromoles.
4) Calculate the proportion between moles of the protein and N-end amino acid residues, is it 1 or 2?
Good luck!
Determine the molecular wt. by gel filtration first to look at the peaks showing activity of respective enzyme ,followed by Native ( PAGE) gel electrophoresis.
Utilize FRET, to determine if the protein is a monomer, or dimer or trimer
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