1. Lecture 10: NMR Applications We may know a tiny bit about NMR theory, but what bioanalytical problems can we adress with NMR? Protein Structures: NMR is one of two main ways of acquiring molecular images of proteins Organic Analysis: Specific organic molecules give characteristic signals in terms of chemical shifts and peak splitting due to J coupling Solid state structures: NMR can be used on solids! But we need to use a special trick, which we’ll talk about Bioanalytical Applications Protein/Protein, Protein/Ligand Interactions: NMR can be used to ‘track down’ binding interfaces Protein dynamics: NMR is the only way to get atomic level information about how proteins move.

2. So You Want To Be An NMR Spectroscopist… If you’re a bioanalytical NMR spectroscopist, here’s the typical runup to an experiment: 1) Grow up your protein with the appropriate label. You’ll either be expressing your protein in bacteria (probably E. coli) or yeast (probably S. Cerevisiae) For 2D NMR: Probably 15 NH 4 Cl or 13 C-  -D-Glucose For 3D NMR: Probably 15 NH 4 Cl and 13 C-  -D-Glucose 2) Purify your labeled protein (probably His 6 or GST tag) 3) Dilute to desired conc. (probably in water, around 1 – 10 mM), add 5-10% D 2 O, 100 uM DSS 4) Make sure the protein is stable under these conditions, place in NMR tube, throw into instrument

3. So You Want To Be An NMR Spectroscopist… 6) Shim the instrument: We have to make the magnetic field perfectly homogenious across the sample or equivalent nuclei will have different spins! The computer can do this for you using the ‘gradient’ approach discussed last time. Shims are extra magnetic coils with their fields pointed in essentially every direction relative to the sample. They can therefore ‘add’ or ‘subtract’ to the big huge magnetic field as needed to even it out.  shim x shim z 5) Set the temperature, check tuning/matching

4. So You Want To Be An NMR Spectroscopist… 7) Calibrate the hard pulse length for 90°: time to 360°/4!! -y z  z  8) You’re ready to go! Load and calibrate the experiment you want to do!! You may have to work out the appropriate power and/or duration of certain soft pulses, depending on the experiment and water suppression scheme. Good tutorial on biological nmr: http://www.nmr.sinica.edu.tw/Cours/Course20040906/NMRExperiments_ LargerMolecule.pdf http://www.nmr.sinica.edu.tw/Cours/Course20040906/NMRExperiments_ LargerMolecule.pdf

5. More on Gradients: DOSY We’ve said that you can destroy magnetization using a gradient pulse. But you can also reconstitute it by using the same pulse with the opposite phase at a later time: Of course, this will only work if the nuclei remain essentially stationary over the course of the wait period. This is the basis for diffusion measurements (DOSY) by NMR!

6. Water Suppression Biological experiments are carried out in water. If we want to see protons from our sample we’re going to need to strongly suppress the water signal. Here are a few ways of doing that: Watergate: Selective ‘soft’ pulses on H 2 O protons 1H1H GzGz Defocus everything Defocus water, refocus not water 3-9-19 Watergate:

7. Water Supression Flipback Watergate: Puts water on z before first gradient pulse Pre-saturation: Lengthy, continuous ‘soft’ excitation of water offset

8. NMR of Peptides Now that we’ve suppressed our water signal, we can take some spectra of peptides in water. If we do a simple water suppression pulse-acquire experiment, we may see something like this:

9. NMR of Peptides: TOCSY That can be useful – we have some idea of what we’re looking at… but which peak corresponds to which specific proton? TOCSY TOCSY tells us which amino acid belongs to which peak

10. NMR of Peptides: NOESY But we still don’t know the amino acid sequence. For that we need to look at ‘through space’ interactions: NOE NOEs are a relaxation effect. As such they are dependent on the correlation time:

11. 1D Saturation Transfer Saturation transfer is a simple technique that can be used to determine if and how something small is binding to something very big. Saturation here is the same as pre- saturation in water suppression. It involves continually hitting a select frequency with a train of soft pulses: Sat. pulse

12. Protein NMR: We Need More Nuclei 1D and H-H coupling experiments are all well and good when you’ve got < 200 protons, but proton signals are not really all that well dispersed. We’re going to need to use couplings between two different nuclei (heteronuclear NMR). Since we’re dealing with proteins our options are most likely 13 C (very expensive!) and 15 N (expensive). The most dispersed signals would be the carbons, but nitrogen is much cheaper! Thus, the most common type of protein NMR spectrum is an HSQC which usually correlates the amide nitrogen with the amide proton. Thus there is one peak per residue (except proline!!)

13. A 13 C Protein Spectrum Here’s a 13 C Spectrum of an SH3 domain (aprox. 70 a.a.):

14. HSQC Spectra Here’s the pulse sequence for the HSQC experiment: And here’s the result: This is ‘the’ HSQC for properly folded Sso Acylphosphatase (104 a.a.) I S x/-x y

15. Detecting and Localizing Ligand Binding Most analytical techniques work hard to tell us ‘if’ something is binding to our protein of interest. NMR not only tells us that, but where! The most common way of measuring this is by ‘ligand titration’ experiments which amount to monitoring the HSQC as a function of ligand concentration. Low Ligand Med Ligand High Ligand

16. Protein Dynamics by NMR: H/D Exchange Once you’ve got an HSQC, you can study slow (minutes to days) conformational dynamics by NMR. To do this, you calibrate your HSQC for speed, ‘buffer exchange’ your protein into 90%+ D 2 O, RUN to the NMR instrument, drop your sample in, quick re-shim and GO!! Here’s the result: No D 2 O 1 st HSQC after D 2 O t = 60 min

17. HDX results Since we know which backbone protons correspond to which signals, we can identify which are more protected:

18. H/H Exchange: CLEANEX CLEANEX is a cool HD exchange technique that uses water protons instead of 2 H! H H H O H H H H Backbone and water protons are exchanging all the time Instead of exciting all protons except water, we only excite water These magnetized protons now exchange onto the protein… And we use that magnetization to transfer to 15 N

19. Results of CLEANEX In order for CLEANEX to work, exchange has to occur faster than the relaxation of protons on the protein. This means mid-to-low milliseconds range:

20. Sequencing Proteins by NMR The HSQC gives us a spectrum in which each amino acid is distinguishable, but doesn’t tell us much about which amino acid they are, and in what order. To do that, we need to extend our analysis into the 13 C plane. 3D NMR!!

21. Sequential Assignment by NMR To do ‘sequential assignments’, we use pairs of J-coupling- based 3D experiments, the most common pair is: C1C1 HNCA C2C2 C C O O N N H H HNCOCA C1C1 C2C2 C C O O N N H H

22. Getting Structural Info: The CSI The Chemical Shift Index (CSI) is a quick way of assessing secondary structure: RESIDUE TYPE HA CA CB CO Ala 4.35 52.5 19.0 177.1 Cys 4.65 58.8 28.6 174.8 Asp 4.76 54.1 40.8 177.2 Glu 4.29 56.7 29.7 176.1 Phe 4.66 57.9 39.3 175.8 Gly 3.97 45.0 - 173.6 His 4.63 55.8 32.0 175.1 Ile 3.95 62.6 37.5 176.8 Lys 4.36 56.7 32.3 176.5 Leu 4.17 55.7 41.9 177.1 Met 4.52 56.6 32.8 175.5 Asn 4.75 53.6 39.0 175.5 Pro 4.44 62.9 31.7 176.0 Gln 4.37 56.2 30.1 176.3 Arg 4.38 56.3 30.3 176.5 Ser 4.50 58.3 62.7 173.7 Thr 4.35 63.1 68.1 175.2 Val 3.95 63.0 31.7 177.1 Trp 4.70 57.8 28.3 175.8 Tyr 4.60 58.6 38.7 175.7 To your observed shifts, give score: +1 if >.7 ppm higher than CSI value -1 if >.7 ppm lower than CSI value 0 if within -.7 to +.7 of CSI value Four shifts in a row at -1 HA and/or +1 CA/CO = minimum for Helix Three shifts in a row at +1 HA and/or -1 CA/CO = minimum for  -strand All other regions are designated random coil

23. Getting Structural Info: NOEs In NMR, the Nuclear Overhauser Effect is the effect that one nucleus has on the relaxation of another. The intensity of this effect is directly related to the proximity of the interacting nuclei: Just like in J coupling, NOE coupled nuclei will experience an oscillating phase at each other’s offsets . ‘is proportional to’ absolute distance between the interacting nuclei correlation function – describes attenuation (or buildup) of the NOE due to the relative motions of the nuclei So the internuclear distance effects the size of the NOE This tells us which nucleus is interacting with which, but a 3D experiment (e.g. HSQC-NOE) is required to distinguish

24. THE HSQC-NOE experiment Here’s the most common NOE-based experiment for structure elucidation: Has the advantage of not requiring double labeling Gives us a set of inter-proton distance contraints We know which amide proton is which and which amide protons are nearby (1.6 – 6 Å).

25. More Structural Info: Angle Restraints A network of NOEs from an HSQC-NOE is a start, but there’s plenty it doesn’t tell us, particularly which way the side chains are pointed. One ‘cheating’ way is to use  /  values consistent with the secondary structures derived from CSI, but there are weak constraints A better way is to do a ‘residual dipolar coupling’ experiment in which the sample is placed in a medium, such as polyacrylamide or phage coat particles, that causes a net alignment with the magnetic field. B0B0

26. Residual Dipolar Couplings Through bond (J) dipolar couplings have well defined frequencies called coupling constants. It is, in fact, by using coupling constants that we pass magnetization around through bonds (such as in a TOCSY). I  J 1,2 The magnitude of the coupling constant depends on the orientation of the interaction with respect to the big huge magnetic field. By measuring the coupling constant (which can only be done in alignment media), we can figure out the bond angle with respect to B 0. difference between aligned and unaligned J angle with respect to B 0

27. Results of Residual Dipolar Couplings Here’s what RDC’s look like. We have to run our HSQC without decoupling. Now we have distance constraints and some bond angles. Combined, these are sufficient to allow us to parameterize a model protein structure. The next step is to plug our distance and angular constraints into a computer program that uses a molecular mechanics force field to find the lowest energy structures that meet the constraints.

28. NMR Structure Results Here’s what an NMR ‘structural ensemble’ looks like: You can then take the average structure or the ‘best’ structure (the one that best fits the constraints) to give you a final structure: Note that, unlike x-ray crystallography, this is a structure from the protein in solution. There are currently 4,448 protein structures in the BioMagResBank database.

29. Dynamics by NMR Since we’re looking at our protein in solution, it should also be moving around roughly like it does in vivo. NMR will allow us to get site specific information about these movements. NMR is also the only method by which motions on virtually all relevant time-scales are observable: Virtually every type of protein motion/activity is covered.

30. Very Fast Motions: T 1 and T 2 We talked about longitudinal (T 1 ) and transverse (T 2 ) relaxation (biophysicists call them R 1 and R 2 ) To make a very long story short, you can get a general description of the conformational freedom for nuclei in a protein by mapping out the spectral density function J(  ) which is directly related to T 1 and T 2. The spectral density function at any particular frequency  is related to the order parameter S via the following relationship: An order parameter of 1 indicates complete restriction of fast timescale motions while S = 0 indicates completely unrestricted motion.  m = correlation time  = correlation time + timescale of bond vibration (~ns)

31. Slower Motions: Relaxation Dispersion NMR Until recently, there was a big hole in the timescale accessible to NMR measurements. And it was centered right on the all important millisecond timescale. The reason is that the equation for the spectral density function becomes underdetermined when an additional term to account for conformational exchange is added. The answer was developed at nearby U of T in the Kay group. They advanced a technique called Carr-Purcell-Meiboom-Gill (CPMG) Relaxation Dispersion NMR.

32. CPMG relaxation dispersion The key to CPMG relaxation dispersion is that the contribution to J(  ) from R ex is suppressed by the application of a train of  pulses. As a consequence, the contribution to J(  ) from R ex alone can be measured and the frequency of the motion causing R ex can be acquired. Chem. Rev. 2006, 106, 3055-3079 A major advantage of CPMG RD is that is sensitive even to low population protein folding intermediates.

33. Solid State NMR Remember we said we can’t see big stuff in solution by NMR because to correlation time is too long and thus T 2 is too fast. Well, what about solids? They aren’t tumbling at all, so they have infinitely long  c and thus an (almost) infinitely fast T 2. (Also recall that, after an initial rise, T 1 goes down with increasing  c ). BUT – this T 2 relaxation is almost all secular, meaning that it is due primarily to dipolar couplings In solution, random tumbling causes these dipolar couplings, which are vector quantities, to cancel each other out Fortunately, some very clever people thought of another way of making this happen…

34. Solid State NMR In solid state NMR, we tilt the sample to the ‘magic’ angle, which is 54.74° relative to B 0.  =54.74° B0B0 And then we spin it around that angle at very high frequency. Thus the name of this type of NMR – ‘Magic Angle Spinning’. To be effective, this spinning has to be close to or above the offset frequency  of the nuclei being observed. We don’t use this too much in bioanalytical chemistry… YET!

35. We’re Done!!! At times, you might have felt like this… But now you’re almost like THIS!