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What Can NMR Do For You? — Part Two

What Can NMR Do For You? -- Part Two

Welcome to part two of “What Can NMR Do For You?”, a three-part series in which we see how you can use simple NMR experiments in your research. In part one, we went over some key points to keep in mind when doing NMR on proteins and DNA, such as sample preparation, and saw how NMR can be used to assess protein folding. In this article, we will see how NMR can be used to determine whether 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.

First, why use NMR to look at protein dimerization? NMR has two main advantages over other techniques. First, NMR can detect weak dimers, with 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 if you’re working with protein domains, which may have weak affinity in isolation and a stronger tendency to dimerize in the context of the whole protein. Second, generating your protein sample and collecting NMR spectra are relatively quick. You don’t have to make two differently tagged proteins like you do for a pull-down, which may interfere with binding, and the spectra themselves take minutes to collect.

To understand how you can measure protein dimerization with NMR, we need to go into a little bit of how NMR works. 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 a certain size, though the size limit of NMR is continuously increasing. If you can measure the rate at which the signal decays, you can get an idea of the size of your molecule. Luckily, you can!

Let’s take another look at a 1D NMR spectrum of a protein:

What Can NMR Do For You? -- Part Two

Let’s ignore all the complexity for now, and just look at the area underneath all of the peaks in the amide region (dotted blue line): we’ll call that the signal intensity. Now, 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 rate of decay. It might look something like this:

What Can NMR Do For You? -- Part Two

(Note: These are not real data. No real data ever look this good.)

In NMR, there are two main rates of interest: T1 and T2. They give you slightly different information, but the basic idea is the same: hit your sample with a certain frequency of radiation, wait a little bit, and then collect a spectrum.

So, what does all this have to do with dimerization? Well, if you know T1 and T2, you can calculate what’s known as the rotational correlation coefficient of your protein, which (drumroll please) is inversely proportional to the size of the protein. So, if you know you have an 8 KD protein, and the rotational correlation coefficient is more consistent with that of a 16 KD protein, you probably have a dimer. I’m not going to bore you with the equations, but here’s a nice article that goes through the details of monitoring dimerization by 1D NMR. If you’re feeling especially industrious, you could even generate the rotational correlation time as a function of protein concentration and get a sense of how weak or strong the dimerization is.

So, next time you’re thinking of measuring dimerization, consider NMR. 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.

What do you think? How do you monitor protein dimerization? Leave your ideas in the comments below.


  1. colinhockings on October 30, 2012 at 12:25 am

    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?

    • Jennifer Cable on November 30, 2012 at 4:31 pm

      Hi Colin,

      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…

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