The Basics of NMR Part 2: NMR and Protein Dimerization
Welcome to part two of “The Basics of NMR?” a three-part series in which we see how you can use simple NMR experiments in your research.
In part one, we learned how NMR can be used to assess protein folding.
In this article, we look at NMR and protein dimerization. I’ll explain how to use NMR to check if a protein is a dimer in solution.
All the key points from part one still apply, and, because we are keeping this simple, we will be focusing on 1D experiments that do not require isotopic labeling of your sample.
Why Study Quaternary Structure Using NMR?
NMR has two main advantages over other techniques.
1. NMR Detects Weak Dimers
The first reason is that NMR can detect weak dimers with a mM-range affinity.
Depending on your system, that may or may not be an advantage since the biological relevancy of such interactions is questionable.
However, this approach may be useful when working with individual protein domains, which may have weaker affinity in isolation than they do when part of the full protein.
2. NMR Is Relatively Quick
The second reason—generating your protein sample and collecting NMR spectra are relatively quick.
The spectra take minutes to collect.
Plus, you don’t have to make two differently tagged proteins as you do for a pull-down assay. So you never end up in a scenario where the tags interfere with binding.
Measuring Protein Dimerization Using NMR
Before we get into details, we need to discuss the basic principles of NMR.
How Does NMR Work?
NMR is like any other type of spectroscopy. You hit your sample with a certain frequency of radiation, the sample absorbs that radiation and produces a signal, and the signal decays over time.
In NMR, the bigger the molecule, the quicker the signal decays. This is why it’s difficult to study molecules above ~30-35 kDa.
If you can measure the rate at which the signal decays, you can estimate the size of the sample molecule. Luckily for us, you can!
Take a look at the 1D proton NMR spectrum of a protein below in Figure 1.
Let’s ignore all the complexity for now and just look at the area underneath the peaks in the amide region (dotted blue line).
This area represents signal intensity.
What if you waited a short time (we’re talking milliseconds here) between irradiating the sample and collecting the spectrum?
Well, you would see the same spectrum, but because the signal is decaying over time, the signal intensity would be a little bit weaker.
If you collect several spectra as a function of different waiting times (also known as delay times), you can fit those data to an exponential curve and graph the decay rate. It might look something like Figure 2.
Note: These are not real data. No real data ever look this good.
In NMR, there are two main rates of interest:
These two rates give you slightly different information, but the basic idea is to hit your sample with a certain frequency of radiation, wait a little bit, and then collect a spectrum.
NMR and Quaternary Structure
What does all this have to do with dimerization?
It’s really simple!
If you know T1 and T2, you can calculate what’s known as the rotational correlation coefficient of your protein.
This (drumroll, please) is inversely proportional to the size of the protein.
For example, if you know your sample protein is 8 kDa, but the rotational correlation coefficient is consistent with a 16 kDa protein, it’s a dimer.
If you’re feeling especially industrious, you can even generate the rotational correlation time as a function of protein concentration and get a sense of how weak or strong the dimerization is.
I wont to bore you with the equations but check out this paper by Aramini et al. for all the details of monitoring dimerization by 1D NMR. 
NMR and Quaternary Structure Summarized
So, now you know all about NMR and protein dimerization!
Also, if you’ve got an NMR enthusiast nearby, ask him or her what extra information you could get from isotopically labeling your protein. You may just be able to determine the dimerization interface, test conditions that disrupt or promote dimerization, or see if other molecules bind to your protein!
Did you enjoy this article? Be sure to leave your ideas, questions, and comments below.
Also, check out part 3 of this NMR series, where we take a look at Multidimensional NMR.
Originally published October 2012. Reviewed and republished on May 2022.
Aramini JM et al. (2011) Dimer interface of the effector domain of non-structural protein 1 from influenza A virus: an interface with multiple functions. J Biol Chem 286:26050–60
Thanks Jenn, very nice article.
I have a protein of around 25kDa which purifies as an oligomer – we think a tetramer. Would a 100kDa complex be too large for this sort of analysis?
100 kDa isn’t impossible to analyze by NMR, but I’d say that unless you have some help from a heavy-duty NMR lab, it’s probably going to give you more of a headache than help. The spectrum I show above in the article is of a 25 kDa protein. I’d imagine, that with a 100 kDa complex, the peaks would become so broad that they would just look like one big lump. Although, since you’d really only be interested in the area under all the peaks, that may not matter. I think the main concern here would be whether you’d be able to get enough signal to noise.
It might be interesting to collect NMR spectra at different concentrations, for example, concentrations at which the monomer or dimer might dominate, as long as those concentrations aren’t so low that you can’t detect anything.
I guess what I’m saying is, it’s probably not impossible, but it’s not ideal. Also, if you’re just doing a 1D, you’ll know within minutes if it’s worth pursuing or not, so, maybe…