Nuclear Magnetic Resonance Spectroscopy (NMR)

An Introduction to NMR

Nuclear Magnetic Resonance spectroscopy (NMR) is a technique that can be used for spectral elucidation of molecules. It is typically used to identify the carbon – hydrogen backbone of an organic structure. The first noted experimental observation of NMR was accomplished just after World War II in 1946 by two individual research groups at Harvard by Purcell and at Stanford by Bloch, whom both received the Nobel Prize in Physics in 1952 for their accomplishments¹. In fact, NMR has been such a valued technique in the scientific community that advancements in the field have been awarded the Nobel Prize three separate times in 1952, 1991, and 2002².

NMR works based off the spinning of atomic particles. The atomic nucleus (¹H, ¹³C, ¹⁵N, etc.) is a spinning charged particle that generates a magnetic field. Without an external force these particles spin at random, however, by applying an external magnetic field the particles spins can be aligned in a single direction either with the magnetic field or against. article with a magnetic field and measuring the magnetic momentum. By sending a magnetic pulse perpendicular to the magnetic field, it causes the magnetic momentum to initially align with the magnetic pulse then relax back to align with the magnetic field. The time it takes for the spin to relax to its original position in the magnetic field can be measured then analyzed through Fourier Transform. Depending on the electronics of the individual atom, and spatial arrangement the signal for the nuclei will produce different signals which allows for the elucidation of small molecules by using a variety of 1-D and 2-D NMR techniques.

Protein NMR

The use of NMR in polymer and protein analysis has been of a more recent endeavor, as the overlapping of monomer units tends to allow for broadening of the ¹H NMR peaks because the repeat units while chemically similar, still have different chemical environments causing a slight peak shift. This slight shift when summed over all similar nuclei will cause overlap to occur desensitizing the NMR spectra (Figure 2). Additionally, there is the issue of the protein being folded or unfolded. If the protein is folded there will be additional non-covalent interactions that
will increase the overlapping of NMR signals which can be seen in figure 2.

NMR is one of two methods to study the three-dimensional structure of proteins. Because of this, new techniques have been developed to analyze proteins using NMR. Typically, ¹H, ¹³C, and ¹⁵N are the isotopes used to analyzed proteins due to their abundance in the desired molecules. Additionally, the use of 2-D NMR techniques correlating two sets of nuclei, or with triple resonance experiments  that were more recently developed in the early 90’s³. Triple resonance experiments are a type of NMR correlation between ¹H, ¹³C, and ¹⁵N which produces a three-dimensional plot that enables an individual to determine the back bone of the protein through the peptide bond linkage (Figure 2)⁴. While this can be a powerful tool in the elucidation of proteins, the chemical shift assignments remain a bottleneck for large proteins due to the faster relaxation times and the degeneracy of the ¹³C_α nuclei⁵.

Dynamic Nuclear Polarization NMR

More recent developments regarding the analysis of proteins by nuclear magnetic resonance is using dynamic nuclear polarization (DNP)⁶. DNP is where spin polarization is transferred from electrons to nuclei  resulting in the alignment of the nuclear spin are aligned with the electron spins. In DNP the samples are doped with a stable paramagnetic polarizing agent and irradiated with microwaves to transfer the high polarization in the electron spin reservoir to the nuclei of interest⁶. This enhances the NMR signals by transferring the large polarization of electron to nearby nuclei via microwave irradiation of electron nuclear transitions, making DNP a vital too in structural and mechanistic studies of biologically relevant molecules⁷.

The use of DNP as an enhancement to NMR has been of utter interest to protein NMR chemists recently. With this agenda in mind, several different enhancements to DNP-NMR have been achieved. Hilty was able to perform real time analysis of folding upon binding of a disordered protein by using dissolution DNP-NMR⁸. This allows for signal enhancements by dissolution DNP after rapid mixing allowing nuclei with lower sensitivity such as ¹³C. Direct ¹³C hyperpolarization of a protein allowed for a large chemical shift dispersion of the nuclei to assess structural changes during real time folding. Hilty was also able to manipulate hyperpolarized binding pockets using the nuclear Overhauser effect for determination of competitive ligand binding⁹. This allowed for DNP to polarize the ³H spins of one of two protein ligands by transferring of a fraction of the polarization of one ligand through the protein binding pocket to the second ligand. Garcia developed a method to reduce the strength of the magnetic field that DNP was able to work by using L-ban electron spin frequencies, allowing the magnetic field to decrease from 0.35T to 0.04T ¹⁰. Sergeyev was able to increase the efficiency of DNP by incorporating non-uniform sampling  to achieve time-effective collection of multidimensional data and by using fast magic-angle spinning (MAS) to provide optimal sensitivity and resolutions¹¹. This allowed for the efficient assignment and NMR analysis of an intact Pf-1 virus using sequential side-chain correlations and DNP sensitization. Rosay also utilized MAS in combination with solid state NMR (SSNMR) to determine the structures of molecules in states that are otherwise not accessible by X-ray or solution NMR experiments¹².

With these advancements, DNP has made the elucidations of proteins more efficient and accessible. Alexandrescu was able to make ¹H NMR assignments of aromatic residues in bovine, human and guinea pig variants of α-lactalbumin by using photochemically induced DNP which enabled assignments to be made in otherwise ambigous areas of the protein structure¹³. Additionally, Williams also used photochemically induced DNP to study calcium binding proteins such as histidine microenvironments in α- and β-parvalbumins. Other interactions between proteins and carbohydrates were analyzed by Siebert et. al. by using a combination of DNP NMR, electrospray ionization mass spectroscopy and molecular modelling¹⁴. This brought about more interest in using DNP in conjugation with molecular dynamic (MD) simulations through the work by Sezer, who was able to calculate DNP coupling factors through the synergestic use of MD simulations and modeling the diffusion of hard spheres with spins as applied to nitroxides in water¹⁵. The Griffin group demonstrated that DNP enhanced MAS NMR experiments performed at 100K was able to produce NMR spectra with excellent signal-to-noise ratios and sufficient resolution to observe intermolecular heteronuclear correlation in mixed samples confirming a parallel in register structure in amyloid fibrils¹⁶. The structural analysis of HIV-1 protein assemblies was able to be determined by Gupta et. al. by utilizing DNP based MAS NMR securing structural analysis of multiple well-resolved isoleucine side-chain conformers; unique intermolecular correlation across two CA molecules; and conformationally disorder states that are otherwise not visible at ambient temperatures ¹⁷.  Chappuis reported the combination of dissolution DNP and water-LOGSY (ligand observed via gradient spectroscopy) in view of high-throughput detection of protein ligand interactions¹⁸. The determination of the cholesterol-binding site of eukaryotic membrane proteins in native like lipid membranes was made possible through Elkins by using low temperature DNP¹⁹. Of note is work by Kendra K. Frederick who has applied DNP NMR to investigate the structure of a protein containing and environmentally sensitive folding pathway and an intrinsically disordered region²⁰. More recently her work has focused on combining DNP NMR with segmental and specific labelling to study a yeast prion protein strain by measuring long range interactions to distinguish between to models for the arrangement of monomers in amyloid fibers²¹.

Dynamic nuclear polarization has been shown to be an essential tool in the structural elucidation of proteins, as well as helping to establish structural conformation of protein interactions. Due to is invaluable nature the future of this application will most likely head toward increasing the accessibility of the method. Typically, protein NMR has to be carried out on larger NMR magnetics that have magnetic fields greater than 600MHz, for long periods of time. Because of this analysis of proteins can be quite expensive. DNP helps to decrease the need for such large magnetic fields and has been decreasing overall analysis time. Thus, it is feasible to say that the future of DNP NMR is to make the application more accessible and enable analysis and structural elucidation of proteins on smaller NMR (<400MHz) and to make the experiments last minutes versus days.

References

1. Nageswara Rao, B. D., Nuclear magnetic resonance – An early history. Resonance 2015, 20 (11), 969-985.

2. Marion, D., An Introduction to Biological NMR Spectroscopy. Mol. Cell. Proteomics 2013, 12 (11), 3006-3025.

3. Shang, Z.; Swapna, G. V. T.; Rios, C. B.; Montelione, G. T., Sensitivity Enhancement of Triple-Resonance Protein NMR Spectra by Proton Evolution of Multiple-Quantum Coherences Using a Simultaneous 1H and 13C Constant-Time Evolution Period. J. Am. Chem. Soc. 1997, 119 (39), 9274-9278.

4. Kay, L. E.; Ikura, M.; Tschudin, R.; Bax, A., Three-dimensional triple-resonance NMR Spectroscopy of isotopically enriched proteins. Journal of Magnetic Resonance 2011, 213 (2), 423-441.

5. Quingtao Wei, J. C., Juan Mi, Jiahai Zang, Ke Ruan, Jihui Wu, NMR Backbone Assignments of Large Proteins by Using 13Ca-Only Triple Resonance Experiments. Chem. – Eur. J. 2016, 22, 9556-9564.

6. Ni, Q. Z.; Daviso, E.; Can, T. V.; Markhasin, E.; Jawla, S. K.; Swager, T. M.; Temkin, R. J.; Herzfeld, J.; Griffin, R. G., High Frequency Dynamic Nuclear Polarization. Accounts of Chemical Research 2013, 46 (9), 1933-1941.

7. Maly, T.; Debelouchina, G. T.; Bajaj, V. S.; Hu, K.-N.; Joo, C.-G.; Mak-Jurkauskas, M. L.; Sirigiri, J. R.; van der Wel, P. C. A.; Herzfeld, J.; Temkin, R. J.; Griffin, R. G., Dynamic nuclear polarization at high magnetic fields. The Journal of chemical physics 2008, 128 (5), 052211-052211.

8. Ragavan, M.; Iconaru, L. I.; Park, C.-G.; Kriwacki, R. W.; Hilty, C., Real-Time Analysis of Folding upon Binding of a Disordered Protein by using Dissolution DNP NMR Spectroscopy. Angew. Chem., Int. Ed. 2017, 56 (25), 7070-7073.

9. Lee, Y.; Zeng, H.; Mazur, A.; Wegstroth, M.; Carlomagno, T.; Reese, M.; Lee, D.; Becker, S.; Griesinger, C.; Hilty, C., Hyperpolarized Binding Pocket Nuclear Overhauser Effect for Determination of Competitive Ligand Binding. Angew. Chem., Int. Ed. 2012, 51 (21), 5179-5182, S5179/1-S5179/16.

10. Garcia, S.; Walton, J. H.; Armstrong, B.; Han, S.; McCarthy, M. J., L-band Overhauser dynamic nuclear polarization. J. Magn. Reson. 2010, 203 (1), 138-143.

11. Sergeyev, I. V.; Itin, B.; Rogawski, R.; Day, L. A.; McDermott, A. E., Efficient assignment and NMR analysis of an intact virus using sequential side-chain correlations and DNP sensitization. Proc. Natl. Acad. Sci. U. S. A. 2017, 114 (20), 5171-5176.

12. Rosay, M.; Lansing, J. C.; Haddad, K. C.; Bachovchin, W. W.; Herzfeld, J.; Temkin, R. J.; Griffin, R. G., High-frequency dynamic nuclear polarization in MAS spectra of membrane and soluble proteins. J. Am. Chem. Soc. 2003, 125 (45), 13626-13627.

13. Alexandrescu, A. T.; Broadhurst, R. W.; Wormald, C.; Chyan, C. L.; Baum, J.; Dobson, C. M., Proton NMR assignments and local environments of aromatic residues in bovine, human and guinea pig variants of α-lactalbumin. Eur. J. Biochem. 1992, 210 (3), 699-709.

14. Siebert, H.-C.; Lue, S.-Y.; Frank, M.; Kramer, J.; Wechselberger, R.; Joosten, J.; Andre, S.; Rittenhouse-Olson, K.; Roy, R.; von Lieth, C.-W.; Kaptein, R.; Vliegenthart, J. F. G.; Heck, A. J. R.; Gabius, H.-J., Analysis of protein-carbohydrate interaction at the lower size limit of the protein part (15-Mer Peptide) by NMR spectroscopy, electrospray ionization mass spectrometry, and molecular modeling. Biochemistry 2002, 41 (30), 9707-9717.

15. Sezer, D., Rationalizing Overhauser DNP of nitroxide radicals in water through MD simulations. Phys. Chem. Chem. Phys. 2014, 16 (3), 1022-1032.

16. Bayro, M. J.; Debelouchina, G. T.; Eddy, M. T.; Birkett, N. R.; MacPhee, C. E.; Rosay, M.; Maas, W. E.; Dobson, C. M.; Griffin, R. G., Intermolecular Structure Determination of Amyloid Fibrils with Magic-Angle Spinning and Dynamic Nuclear Polarization NMR. J. Am. Chem. Soc. 2011, 133 (35), 13967-13974.

17. Gupta, R.; Lu, M.; Hou, G.; Caporini, M. A.; Rosay, M.; Maas, W.; Struppe, J.; Suiter, C.; Ahn, J.; Byeon, I.-J. L.; Franks, W. T.; Orwick-Rydmark, M.; Bertarello, A.; Oschkinat, H.; Lesage, A.; Pintacuda, G.; Gronenborn, A. M.; Polenova, T., Dynamic Nuclear Polarization Enhanced MAS NMR Spectroscopy for Structural Analysis of HIV-1 Protein Assemblies. J. Phys. Chem. B 2016, 120 (2), 329-339.

18. Chappuis, Q.; Milani, J.; Vuichoud, B.; Bornet, A.; Gossert, A. D.; Bodenhausen, G.; Jannin, S., Hyperpolarized Water to Study Protein-Ligand Interactions. J. Phys. Chem. Lett. 2015, 6 (9), 1674-1678.

19. Elkins, M. R.; Sergeyev, I. V.; Hong, M., Determining Cholesterol Binding to Membrane Proteins by Cholesterol 13C Labeling in Yeast and Dynamic Nuclear Polarization NMR. J. Am. Chem. Soc. 2018, 140 (45), 15437-15449.

20. Frederick, K. K.; Michaelis, V. K.; Corzilius, B.; Ong, T.-C.; Jacavone, A. C.; Griffin, R. G.; Lindquist, S., Sensitivity-Enhanced NMR Reveals Alterations in Protein Structure by Cellular Milieus. Cell (Cambridge, MA, U. S.) 2015, 163 (3), 620-628.

21. Frederick, K. K.; Michaelis, V. K.; Caporini, M. A.; Andreas, L. B.; Debelouchina, G. T.; Griffin, R. G.; Lindquist, S., Combining DNP NMR with segmental and specific labeling to study a yeast prion protein strain that is not parallel in-register. Proc. Natl. Acad. Sci. U. S. A. 2017, 114 (14), 3642-3647.

22. Figure 1 was photographed by the author. Figure 2 was taken from “Introdution to Protein NMR Spectrocopy” by James Chou file:///C:/Users/HP/Downloads/method2.pdf . Figure 3 was taken from https://en.wikipedia.org/wiki/Triple-resonance_nuclear_magnetic_resonance_spectroscopy .