If other techniques are available, I generally don't recommend the use of cross-linkers  because the process is funadamentally stochastic. Unless you have a very specific cross-linker (for example, attaching to cysteines near the dimerization site) you will get a mixture of dimers and higher order oligomers. This results in false positives at high concentration and false negatives at lower concentrations.

Ion mobility mass spectrometry (IM-MS) requires the dimer complex to be stable in the gas phase. Although it is quick and does give you additional information beyond some of the simple techniques, it would not be my first choice just to see if it is a dimer or not.

The simple techniques are actually the best here. As pointed out by other posters, a native gel or size exclusion column will likely give you the correct answer. Keep in mind, though, that native gels are generally not run in strictly physiological conditions. The pH is adjusted away from neutral  to give the protein enough charge to migrate in the gel .Blue native gels may also mask charges important for complex formation. In contrast, size exclusion columns will give the correct answer in most situations.

For weaker complexes that may fall apart with separation based techniques, DLS ,diffusion NMR, or ITC may help and give you additional information as well (energetics with ITC and residue specific information if you have an assigned spectrum with NMR).

Edit: As Michal Jakob and Scott Kalbach suggest, analytical ultracentrifugation is one of the most accurate methods for measuring monomer/oligomer equilibrium. Unlike all the other techniques except cross-linking, it can also (with difficulty) be used for membrane proteins by adjusting the density of the solvent with D2O or sucrose.

Gel filtration could be the most straight forward way.

analytical gel filtration chromatography

it depends on what you have, if your protein is in a good amount (generally recombinant version) you can go for size exclusion chromatography coupled ligh scattering measurment (where you get the exact molecular weight of your protein). if your protein is not reocombinantly expressed, then you can analyse them through co-IP methods by expressing with 2 different tags.

If you have some time you might try some old fashioned methods like cross-linking or analytical ultracentrifugation.

I would also go for cross-linking and gel analysis.

Besides the methods already mentioned (e.g. gel filtration chromatography, ultracentrifugation, light scattering) you may perform thermal or chemical denaturations. For a monomeric protein the mid transition temperature Tm or the mid denaturant concentration Cm do not depend on protein concentration. However, for an oligomeric protein they depend on protein concentration. That is, increasing the protein concentration will increase the apparent stability of the protein against thermal or chemical denaturation; the intrinsic Gibbs energy difference between the native and the unfolded states is concentration independent though. Therefore, performing two denaturation experiments at two significantly different protein concentrations (e.g. 2 and 20 micromolar), if the Tm or the Cm are different, then the protein is not monomeric in the native state. Of course, I am assuming the unfolded state is not an oligomer, which is usually the case. The denaturation can be performed using spectroscopic techniques or differential scanning calorimetry. In addition, calorimetry allows determining the calorimetric unfolding enthalpy dHcal and the van't Hoff unfolding enthalpy dHvh. If dHvh is larger than dHcal, the native state of the protein is oligomeric. The larger the difference, the higher the stoichiometry of the oligomer.

Important note: this two methods will let you know if the native state of the protein is monomeric or not, but no information about the stoichiometry of the oligomer can be obtained (that is, whether it is a dimer, a trimer, a tetramer,...).

An enzymatic assay (substrate concentration vs. initial rate) could help. If you find that the curve you get has a sigmoidal behavior, at least it tells you that your protein is not a monomer.

dear Silvia,

what if my protein is not an enzyme? and how we can interpret the sigmoidal behaviour with its state? please explain.

Electrophoresis samples of untreated and after treatment with mercaptoethanol, if the two results are consistent, basically that is a monomer, if appears several bands, there may be multimeric.

My experiment idea is to design a mutant protein, this mutant protein does not have function, but the range of mutations must be very small, can not affect the protein folding and quaternary structure formation, the best is point mutations and can't mutate cysteine(because cysteine ​​is often an important group of multimeric), Load this mutant protein and normal protein in the same expression vector, or with two vectors respectively (requires higher expression of mutant proteins compared to normal protein) introduced into a cell for expression, if the protein is a catamaran or multimeric, then there must be a most multimeric mutant embedded, such proteins will not have function;if the protein is a monomeric form, then the expression of normal protein can complete the normal function. This idea actually often used for the analysis of transcription factor :Dominant Negative Mutant analysis.

I agree with Guanfan Mao.The simplest way of finding it out is to run the protein sample in both SDS PAGE and Native PAGE and see the band pattern. If there is only one band at the same size in both the gels,the protein is a monomer and if there is only one band in native gel and a different band pattern in SDS gel,it could be a polymer.

Guanfan Mao and Mallu Abhiram Charan Tej, I think this is a good an idea. I will try to do this! Thanks to all answers!

thermal denaturing and SDS-PAGE provided me with conformation of a dimeric structure of a protein i was looking at, although as an undergraduate i was only following the guidance of others.

Try firstly native polyacrilamide gel electrophoresis and then SDS-polyacrilamide gel electrophoresis with a reductor agent as ditiotreitol or beta-mercaptoethanol in sample buffer and onto the gel.

Run SDS-PAGE under reducing and non-reducing conditions in the same gel, you will surely get some information which will help you.

Why not in separate gels? sometimes it is necessary to add the reductor agent to the gel not only to the sample.

It can be done in two seperate gels also but it will be better to compare the samples in same gel.

Small-angle X-ray scattering (SAXS) would give you the most accurate answer, especially if you have monomer-dimer equilibrium in solution.

The Gold Standard to analyze proteins higher order assemblies is analytical ultracentrifugation or Right Angle Light Scattering coupled in line with analytical gelfiltration. If you have the possibility do both!

You mean the Gold Standard at Max Delbrück Centrum für Molekulare Medizin, I guess :-)

... of course only as we first would have to travel from Berlin to Hamburg to do SAXS dear Salam (-:

By the way: SAXS needs pure protein species, doesn't it!? If you have a fast exchanging equilibrium between monomer:dimer:tetramer i.e. scattering is too inaccurate to determine anything with SAXS?? Please correct me if I am wrong!

With AUC and/or RALS you can determine such equilibria quite easy.

Actually it is not a problem, have a look at the paper we published in 2011, Söderberg et all, JMB (data collected in Hamburg :-)). We analysed iron-induced oligomerisation of a protein called frataxin, which makes many different oligomers, depending on iron concentration in solution. The analysis in such a case assumes of course that you have an X-ray structure to be used for modeling the SAXS data, otherwise it could be problematic, as you say. But if it is just monomer-dimer equilibrium, it should be rather straightforward.

Dynamic light scattering to determine the size of a given particle is a possibility.

Aside from the suggestion to run native page versus SDS-page described above I have used size exclusion chromatography (SEC) extensively to determine if a protein is present as a monomer/dimer etc. Transfer of native gels for western blotting can be tricky, so doing SEC and then SDS-PAGE/western blotting of the resulting fractions can be an easy way to determine the size of any complexes the protein might be in. You can use a fancy machine and an expensive column (we do this alot so it makes sense for us to invest in the HPLC apparatus) or you can do it quickly and easily by hand using a cheap small column or even a 10 mL syringe with the plunger removed and the bottom hole stuffed with glass wool. Another option is to treat a relatively dilute sample of lysate with a crosslinking agent such as dimethyl suberimidate or even plain gluteraldehyde, then do a regular SDS-PAGE gel and western blot, you will see a series of bands of varying sizes depending on what the protein of interest is crosslinked to. A dimer would be indicated by a band exactly twice the size of the momomeric protein. Comparing mercaptoethanol-treated to untreated samples will only determine if the dimer is stabilised by disulfide bonds, and will not distinguish if the dimer is non-covalent. Details on the use of SEC and native/non-native gels to determine if a protein is dimeric etc. are available in our publication "Parameters for effective in vitro production of zinc finger nucleic acid-binding proteins. Belak et al. Biotech. App. Biochem. 58(3)166-174 2011"

compare SDS PAGE and gel permeation chromatography results, u will get a clear cut picture.

Compare the results of SDS-PAGE and Analytical GPC; It will tell you the nature of your protein, very simple!!

I agree with comments from Kumar and Nduko. It is more simple and does not need any expensive tools for analysis.

Perform size-exclusion (also called gel-filtration) chromatography of your protein and compare actual elution volume with the theoretically expected (based on the protein's molecular weight). I used that technique before to study protein dimerization (I attached paper; see Discussion section on the page 4).

Article Dimerization effect of sucrose octasulfate on rat FGF1

Above answers are good. Is the protein an enzyme? A good functional test would be to check if the enzyme activity is associated with dimer -not monomer. Hope this helps!

Ideally use several methods and contrast results, first apply the protein to SDS-PAGE, and then to NATIVE-PAGE and / or gel filtration chromatography. A very useful method is dynamic light scattering (DLS), which can vary ionic strength, pH, protein concentration, ion type, temperature, etc.., And shows you how assembly varies, you can determine particle size and hydrodynamic radius.

I assume you know the mass of your protein and you want to know if it is either monomeric or multimeric in structure. Is that it?

You can try ESI MS if you have a suitable mass spectrometer in your lab. ESI is delicate enough to preserve the bonds between the monomers. You could try nanoESI. You should be able to obtain the envelope of the dimer and the one of the monomer. By the separation of the isotopic peaks in each envelope you can establish their charge state and verify if the m/z make sense for the mass of your protein. Then you can try things that would lead to the break up of the dimer into monomers and observe if the spectrum reflects that.

Every method has its detection limit. There are a lot ways. Size Exclusion, native PAGE, cross-linking, Laser light scattering, analytical ultracentrifugation, diffusion constant measurement(solution NMR).......

Estimate the mol size using gel exclusion chromatography and then compare the results with SDS page to confirm whether the protein is a dimer or not

I agree with comments from Kumar and Nduko too.

Dear Fernanda

If your protein is pure you can use the following comments in addition to PAGE:

You must prepare two samples that one of those contains 6 M urea, 2 M thiourea, SDS and 2ME.

Then you should to run simple gel filtration column chromatography (e.g. G100) to compare elution pattern.

If one peak seen at two different sample, it means that your protein is monomer

If one peak seen in 1st sample and two peaks seen sample contain urea, thiourea and ..., this mean that your protein is dimmer.

Cheers

Ok, let me add my two cents, here as well. I am not an experimentalist and remain exclusively in computation, but I think this idea of whether a protein is monomeric or not is a wonderful fundamental question nowadays.

In the strict wet-lab setting, I have heard from my experimental collaborators, that chemical linkers can be used to see whether your protein has the capabilty to form multimeric assemblies. One can then measure the percentage of dimers, trimer, tetramers, and so on, if they exist. There may be specific limitations to how high one can go in monomeric number, but at least one can see whether dimers to trimers exist or not. Also, one can do this in-vivo or in-vitro to see whether there are any differences. I apologize for not recalling the proper names for these techniques, but, as I said, I am not a wet-lab person.

From a computation point of view, if you have some idea of what the structure of your unit is (or you can estimate it through homology, if you are lucky), then you can use off-the-shelf docking software to at least see whether there are credible dimers that your protein can form. Most software out there are limited to dimers. There is one, MultiZerDock, that can do multimers (CombDock, can do, too, but it produces sometimes less than credible structures). In any case, you could use these computational tools not to be guided by them, but to see whether they are adding some structural information on top of the experiments you are performing. For instance, if your wet-ab investigation points to the fact that this protein may form dimers, than the software can give you some insight onto the structure and actual interaction interface. Then, you can run with this for more studies :-)

If other techniques are available, I generally don't recommend the use of cross-linkers  because the process is funadamentally stochastic. Unless you have a very specific cross-linker (for example, attaching to cysteines near the dimerization site) you will get a mixture of dimers and higher order oligomers. This results in false positives at high concentration and false negatives at lower concentrations.

Ion mobility mass spectrometry (IM-MS) requires the dimer complex to be stable in the gas phase. Although it is quick and does give you additional information beyond some of the simple techniques, it would not be my first choice just to see if it is a dimer or not.

The simple techniques are actually the best here. As pointed out by other posters, a native gel or size exclusion column will likely give you the correct answer. Keep in mind, though, that native gels are generally not run in strictly physiological conditions. The pH is adjusted away from neutral  to give the protein enough charge to migrate in the gel .Blue native gels may also mask charges important for complex formation. In contrast, size exclusion columns will give the correct answer in most situations.

For weaker complexes that may fall apart with separation based techniques, DLS ,diffusion NMR, or ITC may help and give you additional information as well (energetics with ITC and residue specific information if you have an assigned spectrum with NMR).

Edit: As Michal Jakob and Scott Kalbach suggest, analytical ultracentrifugation is one of the most accurate methods for measuring monomer/oligomer equilibrium. Unlike all the other techniques except cross-linking, it can also (with difficulty) be used for membrane proteins by adjusting the density of the solvent with D2O or sucrose.

Is there a reason you suspect that it is not a monomer? Perhaps you could give some more information. Did you observe a band on gel that did not correspond to the monomer size? Anyway, if so, you could also try to express two differently tagged versions (let's say HA- and FLAG) of your protein in cells, (or in E. coli and do it in vitro), and pull down on the one tag (HA), and see if you pull down the other on Westernblot (FLAG). That is assuming that your interaction is strong enough (but if you have this suspicion because of a gel size band it will be). In any case, if it it does function as a dimer, I would suspect you will pull it down. You will need to have the right controls, and % input to be able to make any statements. Alterbatively, you can crosslink the two tagged proteins (should a normal co-IP not work). At least it will tell you whether or not they are interacting (stoichiometry you won't be able to tell). Hope you have some useful approaches here. A gelfiltration column will also tell you something. Most likely, you will always also get a monomer peak in addition to your dimer or oligomer peaks. While there are some caveats with such a column, it would be a good try to... just include the proper size markers. If you need any help, let us know.

We like x-ray scattering for these experiments. As a general rule, it’s worth the time investment to check the assembly state of your macromolecule under multiple conditions and especially under near physiological conditions. Small proteins may be monomers but are more often than not multimers. In a 50-protein test set, employing small angle x-ray scattering (SAXS) of 20ul volumes of ~2ug/ul solutions showed that 60% were multimeric (see GL Hura et al. Robust, high-throughput solution structural analyses by small angle X-ray scattering (SAXS) Nature Methods 6:606-612, 2009). So your protein may have one or more assembly states that should be tested with activity. At the SIBLYS beamline (http://bl1231.als.lbl.gov) , there is a mail-in program, so users can send 96-well plates and get the data back (J Appl Crystallogr. 46,1-13, 2013). It takes 12 hours to run four 96-well plates. SAXS gives direct measurements of mass in solution so defines the assembly state under as many conditions as you might wish to examine, such as with and without ligands, substrates, or inhibitors (Nature 496, 477-81, 2013).

I would consider AUC. If the dimer is in fact forming, then the observed molecular will be twice that of the monomer. This would be a simple fit being that there is only one protein present. This has been the most successful way I have determined that my trimers are forming in my mimic systems.

I would use a Blue-Native PAGE. With this you can easily determine what species you have in solution.

If protein showed single band at 20 kda in both the non-reducing and reducing as well, but showed different mass peak exactly in a manner of like 10kda ,20 kda and 40 kda in MS spectrum. What could be the best possible answer for this ? is protein under going oligomerisation or getting degraded ? please do suggest me

Simple SDS-PAGE  and gel filtration with or without 8M urea treatment should give an idea whether the protein is a monomer or a dimer.                                                  

Hi everybody, you all did mentioned a quite exciting ways, anyhow, you can:

1. run gel electrophoresis >> m.wt

2. urea treatment >> m.wt

3. Beta-mercaptoethanol >> m.wt (if it is thesame as above, then it's monomeric, if different m.wts then it's polymeric...)

to rule out how many subunits there is,, due purification of the resultant m.wts,, then peptide bond map and amino acid sequencing ..

I guess this would be good ;)

Simply, you can run your protein on SDS-PAGE under both reducing condition (with B-mercaptoethanol) and non- reducing condition (without adding B-mercaptoethanol). if you get the same bands in both conditions that means  your protein is a monomer, but if you get higher bands in the non- reducing condition that means it's dimer or trimes depending on the molecular weight. Good luck

SEC-MALS (Size Exclusion Chromatography-MultiAngle Light Scattering) 

SEC-MALS uses multi-angle light scattering to characterize molar mass, molecular weight and size of proteins, polymers, nanoparticles and more.

but if I want want to know that my protein is homodimer or not only by chromatography . then how to proceed

How to differentiate homodimer and heterodimer from the protein complex  

Measurement of hydrodynamic radius (Rh) should provide an indication of oligomeric state. As you're looking at radius, a dimer will not neccessarily have twice the Rh as a monomer, but the difference should be sufficient to make an educated guess.

With this in mind you have a few different options.

Size Exclusion Chromatography - can be used to resolve multiple species, especially if combined with a MALS. However, transit time in column out of equilibrium can be >30 min, which means dimeric species may appear as monomers if interaction is weak.

Analytical Ultra-Centrifugation - will distinguish all species but can take >24h and data interpretation is hard

Dynamic Light Scattering - can be used to determine Rh quickly, but will bias towards aggregates present in solution.

Microfluidic Diffusional Sizing - determines average Rh using 5 uL sample. Time out of equilibrium is minimal (seconds). Does not determine % monomer, dimer etc, but simply gives average. https://www.fluidicanalytics.com/resources/faq/can-fluidity-one-distinguish-between-monomers-dime/

SEC-MALS.