Protein Structure and Interactions from Nuclear Magnetic Resonance Spectroscopy

Ranajeet Ghose, Graduate Center of the City University of New York, New York, USA

Published online: January 2017

DOI: 10.1002/9780470015902.a0003100.pub3

Abstract

Nuclear magnetic resonance (NMR) spectroscopy is a biophysical technique that facilitates determination of protein structures and interactions in solution. NMR experiments are used to identify spectral signatures corresponding to specific atoms on individual protein residues and determine their order in the primary sequence. Distance, angular and orientational relationships are subsequently measured and utilised in computational protocols to obtain high‐resolution structural models. Structure determination has been aided by advances in biotechnology allowing the generation of samples optimally labelled with NMR‐active isotopes; by improvements in the design of NMR instrumentation to allow spectral signatures to be recorded with high sensitivity; by the development of efficient techniques to manipulate nuclear spins at the quantum level and by the creation of computer algorithms that allow the rapid processing and manipulation of NMR data and the incorporation of novel information available from NMR experiments into strategies to obtain the atomic models of proteins and their complexes.

Key Concepts

  • Solution NMR is a spectroscopic technique that enables the determination of the three‐dimensional structure of proteins and their interactions under near‐physiological conditions.
  • The abundance of chemical and structural information available from NMR spectra derives from the ability to selectively manipulate specific nuclei within the molecules placed in a large static magnetic field by customisable sequences of radio‐frequency pulses.
  • NMR signals report on the variety of chemical environments experienced by nuclei in proteins through unique spectral signatures.
  • The repertoire of information‐rich NMR experiments has been significantly expanded by the ability to enrich biomolecules with NMR‐active nuclei by taking advantage of advances in biotechnology.
  • Nuclear spins in proteins form magnetically coupled networks, with characteristic signatures in NMR spectra that can be manipulated to extract angular, distance and orientational information to enable the generation of three‐dimensional structural models.
  • Changes in the spectral signatures of individual nuclei that make up each component of a complex in the presence of the other component/s provide information about the mode of interaction among the constituents in the assembled state. In some cases, where three‐dimensional structures of individual constituents are available, the spectral changes or perturbations may be utilised in novel computational strategies to obtain three‐dimensional structural models of the complexes.

Keywords: NMR; proteins; three‐dimensional structure; biomolecular interactions; isotope labelling

Figure 1. One‐dimensional to multidimensional Fourier transform nuclear magnetic resonance (NMR). (a) Representation of a free‐induction decay (FID) time‐domain signal that upon Fourier transformation yields the resonance frequencies of all the constituent nuclei, that is a spectrum. (b) A typical 2D experimental scheme includes a preparation period, an evolution period (the delay t1 is incremented in discrete steps) and a detection period (t2) during which the FID is detected. The last two periods usually sandwich a mixing period [e.g. in a NOESY (nuclear Overhauser effect spectroscopy) experiment the mixing period is usually 150–300 ms and allows the dipolar coupling between two protons to enable magnetisation exchange before detection]. Fourier transformation of the detected signal in each of the evolution (indirect) and detection (direct) domains leads to a 2D spectrum with two frequency (chemical shift) domains – δ1 and δ2 (expressed in parts per million, ppm). This approach can be generalised to additional dimensions, for example a 3D spectrum has two indirect evolution periods. The 2D spectrum of a hypothetical molecule containing three nuclei (A, B and C) has a 1D NMR spectrum that is displayed along the diagonal of a 2D spectrum (red, blue and green circles). Correlation of the resonance frequencies (chemical shifts, δA, δB and δC) of the atoms leads to off‐diagonal cross‐peaks each representing a correlation, either through‐bond (COSY or TOCSY experiment) or through‐space (NOESY experiment), between the nuclei. In the example shown here, nucleus C is coupled to both nucleus A, and to nucleus B (as evident from the cyan cross‐peaks at δA,δC and δC,δA and the yellow cross‐peaks at δB,δC and δC,δB); nuclei A and B are not coupled to each other hence there are no cross‐peaks at δA,δB or δB,δA. (c) A homonuclear 1H–1H 2D NOESY spectrum of a protein showing through‐space correlations. (d) 2D 15N, 1H heteronuclear correlation spectrum (HSQC) of a protein displaying the protein amide 15N and attached 1HN chemical shifts. Note that some side chains, for example the NH2 moieties of certain amino acids (e.g. glutamine and asparagine; two cross‐peaks at each 1H chemical shift for the same 15N chemical shift) and the indole NH moiety of tryptophan also yield cross‐peaks in an HSQC spectrum. (e) Advantages of the transition from two to three dimensions can be appreciated by the fact that the red cross‐peak in the 2D spectrum appears to be a single peak but resolves into two separate peaks (light and dark orange) along the third dimension.
Figure 2. Selective labelling of proteins. (a) A 2D H(N)CO (in red, bottom; similar to a conventional 3D HNCO spectrum except that the amide N chemical shift is not recorded) spectrum for a protein selectively 13C′‐labelled only on the phenylalanine residues in a uniformly 15N‐labelled background. This labelling is generated by supplementing the growth medium (M9 with 15NH4Cl, unlabelled glucose) with phenylalanine 13C‐labelled at the carbonyl C′ position. The top panel shows a corresponding 2D H(N)CO spectrum of uniformly 13C, 15N labelled protein (in black). Note that not all amino acid residues can be labelled in this simple way due to substantial cross‐talk between their respective biosynthesis pathways that leads to the label being ‘scrambled’. (b,c) Position‐specific labelling can be achieved by supplementing the growth media with precursors of specific amino acids labelled in a particular manner. This scheme exploits the fact that certain chemical moieties on the precursors are incorporated at defined positions in the side chains of the amino acid residues they generate. (b) Specifically labelled α‐ketobutyric acid results in the incorporation of the label at the δ1 position of isoleucine (red) residues. (c) Specific labels placed on α‐ketoisovaleric acid lead to the transfer of the labels to the δ1/δ2 and γ1/γ2 positions of leucine and valine residues, respectively. All other positions are 12C and 2H. Examples of the resulting 13C,1H HMQC (heteronuclear multiple quantum correlation) spectra of the resultant samples are shown. While 13C,1H HMQC spectra provide similar information as 13C,1H HSQC spectra (discussed in the text), they can be recorded with higher sensitivity when using this particular labelling scheme for methyl groups.
Figure 3. Establishing intra‐ and interresidue correlations in proteins. The 3D HNCACB experiment (a) correlates the chemical shifts of the 13Cα and 13Cβ nuclei of a residue (i) and the 13Cα and 13Cβ nuclei of the previous (i − 1) residue with the chemical shifts of its own (i) amide 15N and 1HN nuclei. The 3D CBCA(CO)NH (b) experiment correlates the chemical shifts of the 15N and 1HN nuclei of a given residue (i) with the 13Cα and 13Cβ shifts of the preceding (i − 1) residue. In this example (c), two sets of HNCACB/CBCA(CO)NH pairs are shown that yield (i − 2)/(i − 1) and (i − 1)/i correlations. The 13Cα and 13Cβ resonances are shown in dark blue and red (signifying their opposite phases, when one is positive in sign, the other is negative) for the HNCACB experiments, respectively, and both are shown in blue for the CBCA(CO)NH experiment. The J‐couplings that allow transfer of magnetisation include 1J(CH) ∼ 140 Hz, 1J(NHN) ∼ 92 Hz, 1J(CC) ∼ 35 Hz, 1J(CC′) ∼ 55 Hz, 1J(NC′) ∼ 15 Hz, 1J(NCα) ∼ 7–11 Hz and 2J(NCα) ∼ 4–9 Hz. The flow of magnetisation is indicated by the red arrows and is bidirectional for the HNCACB experiment (a so‐called out‐and‐back experiment) and unidirectional in the CBCA(CO)NH experiment (a so‐called straight‐through experiment). Each strip plot is extracted from the 3D data set at the plane corresponding to the chemical shift of the amide 15N nucleus indicated at the top of each strip.
Figure 4. Sequence‐specific assignment. Four strip plots showing the sequential connectivity between four successive residues (i − 1)/(i)/(i + 1)/(i + 2) through their 13Cα and 13Cβ resonances, the so‐called HNCACB walk. Note that the (i + 2) residue is a glycine and is therefore missing a 13Cβ resonance.
Figure 5. Characteristic chemical shifts for amino acid residues. The mean (represented by circles) and standard deviations (represented by horizontal error bars) for the backbone and side‐chain 13C (a) and 1H nuclei (b) extracted from the Biological Magnetic Resonance Bank (BMRB) are shown. The cysteine Cβ position has a large standard deviation as it includes both the oxidised (in a disulfide bond) and the reduced (SH) forms of the amino acid.
Figure 6. Examples of the types of constraints obtained from NMR experiments. (a) Karplus curves relating measured three‐bond scalar couplings (3J) to dihedral angles. The black, red and blue curves represent the influences of the backbone dihedral angle φ, the backbone dihedral angle Ψ, the side‐chain dihedral angle χ1 on the 3J(HNHα), 3J(Hα,iNi+1) and 3J(HαHβ2) coupling constants, respectively. The parametric equations representing the relationships are given by 3J(HNHα) = 7.97 cos2(φ − 60°) − 1.26 cos (φ − 60°) + 0.63 Hz; 3J(Hα,iNi+1) = −1.0 cos2(Ψ − 120°) + 0.65 cos (Ψ − 120°) − 0.15 Hz; 3J(HαHβ2) = 9.5 cos2(χ1 − 120°) − 1.6 cos (χ1 − 120°) + 1.8 Hz. A qualitative evaluation of the values of 3J(HNHα) can be used to ascertain whether a particular residue is in a helix or in an extended structure like a β‐sheet given that these values are small in the former (α‐helix, φ = −57°, 3J = 2.84 Hz, 310‐helix, φ = −49°, 3J = 1.88 Hz) and quite large in the latter (parallel β‐sheet, φ = −119°, 3J = 9.86 Hz; antiparallel β‐sheet, φ = −139°, 3J = 8.95 Hz). These equations are only one of several parametrisations of the Karplus curves for a given 3J coupling; in this case 3J(HNHα) is from Vogeli et al. , 3J(Hα,iNi+1) is from Lohr et al. and 3J(HαHβ2) is from Demarco et al. . Other forms of the Karplus curves have also been suggested (Schmidt, ). (b) Measurement of residual dipolar couplings (RDCs) from coupled 15N, 1H HSQC spectra. Two sets of coupled HSQC spectra are shown, in isotropic solution (right), and in a medium that leads to weak alignment of the molecule (left). Residual dipolar couplings (D) shown here are all negative in sign leading to decreased splitting of the resonances (J + D) in the partially oriented state. (c) The cross‐peak (red) obtained in a long‐range 2D H(N)CO experiment that optimises magnetisation transfer based on a three‐bond J coupling ( ) between N and C′ nuclei across a hydrogen bond. The cross‐peak appears at the chemical shifts of the nuclei corresponding to the hydrogen‐bonded 13C′ and 1HN nuclei (right panel). Also shown for comparison is an overlay of the corresponding region of a conventional 2D H(N)CO spectrum (optimised for the one‐bond NC′ coupling, ) that show the resonances (blue) at the chemical shifts of the backbone 13C′(i − 1) and 1HN (i) nuclei, there is no peak corresponding to the hydrogen‐bonded pair in this spectrum. A comparison between these two sets of spectra allows the elucidation of the residues that participate in a hydrogen bond.
Figure 7. A flowchart illustrating the steps in determining protein structures by NMR. Following the preparation of isotope‐labelled samples and data collection on the NMR spectrometer, a variety of constraints (distance, angular and orientational; the last is usually not used in the initial rounds) are used as inputs into a calculation protocol termed molecular dynamics simulated annealing. The simulated annealing calculation uses a target function that models the potential energies of chemical bonds, nonbonded energies such as van der Waals forces and angular relationships, in addition to the experimental constraints. Validation is performed at each step in a cycle to check for violations of experimental constraints by the cluster of structures generated; incorrect assignments (these are most commonly incorrectly assigned NOESY cross‐peaks) are rectified; and further assignments are obtained to extract additional constraints; additional experiments are performed if deemed to be necessary. The iterative cycle continues until all possible experimental data are utilised in the calculation. The final structural ensemble is validated for local and global geometry and structural characteristics. The final step consists of deposition of the atomic coordinates into the PDB and the resonance assignments into the BMRB databases.
Figure 8. Structure refinement. Illustration of the improvement in the root‐mean‐squared deviation (RMSD) in a cluster of structures as more and more experimental restraints are incorporated into the structure calculation protocol during the iterative refinement procedure.Reproduced with permission from Shekhtman et al. 2002 © John Wiley and Sons.
Figure 9. NMR spectra to assess molecular interactions and stability. (a) Representative chemical shift changes in 15N,1H HSQC spectra of a protein in the presence of a ligand (blue). The spectra of the apo (ligand‐free) protein are shown in black. The boxed resonance does not change position in the presence of the ligand suggesting that the corresponding region is not involved in binding. The protein/ligand interaction illustrated here is in the fast exchange regime resulting in the gradual shift of the resonances from the chemical shift position of the free state to that of the bound state with increasing titrant concentration. The region for which resonances show chemical shift changes in the presence of the ligand are mapped onto the structure of the protein revealing the binding site. (b) Gradual chemical shift changes (Δδ) with increasing ligand concentration for the fast exchange regime. The different curves represent titration profiles different resonances. The chemical shift changes or perturbations are usually estimated from a 2D correlation spectrum, for example a 2D 15N,1H HSQC by measuring the changes in heteronuclear (e.g. 15N) and proton positions during the titration course. The Δδ(j) at each (jth) titration point can then be represented as Δδ(j) = [(δH,j − δH,free)2 + K(δX,j –δX,free)2]1/2, where δH,j and δX,j represent the proton and heteronuclear chemical shifts for the jth titration point, respectively; δH,free and δX,free represent the corresponding chemical shifts of the proton and the attached heteronucleus in the ligand‐free state; K is a sensitivity factor that normalises the chemical shift responses of proton and heteronucleus to changes in local environment. Fits (denoted by the straight lines) of the experimental Δδ (j) (circles) against ligand concentration to standard binding isotherms (1‐site binding in this case) yields the apparent binding affinity (KD). (c) Generation of a double tyrosine‐to‐alanine mutant in the core region of a helical protein leads to a significant loss in chemical shift dispersion (red) compared to the wild‐type spectrum (blue) suggesting that the protein unfolds. This represents an example of NMR spectra being used to assess the structural effects of mutations and this approach can be used as a protein quality control step when performing structure‐based mutations to test binding or activity (in enzymes).
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References

Anderson WA and Freeman R (1962) Influence of a second radiofrequency field on high‐resolution nuclear magnetic resonance spectra. Journal of Chemical Physics 37: 85–103.

Arnold JT, Dharmatti SS and Packard ME (1951) Chemical effects on nuclear induction signals from organic compounds. Journal of Chemical Physics 19: 507.

Aue WP, Bartholdi E and Ernst RR (1976) Two‐dimensional spectroscopy. Application to nuclear magnetic resonance. Journal of Chemical Physics 64: 2229–2246.

Baldisseri DM, Pelton JG, Sparks SW and Torchia DA (1991) Complete 1H and 13C assignment of Lys and Leu sidechains of staphylococcal nuclease using HCCH‐COSY and HCCH‐TOCSY 3D NMR spectroscopy. FEBS Letters 281: 33–38.

Bardiaux B, Malliavin T and Nilges M (2012) ARIA for solution and solid‐state NMR. Methods in Molecular Biology 831: 453–483.

Bloch F (1946) Nuclear induction. Physical Review 70: 460–474.

Bloch F, Hansen WW and Packard M (1946) Nuclear induction. Physical Review 69: 37.

Bodenhausen G and Freeman R (1977) Correlation of proton and carbon‐13 NMR spectra by heteronuclear two‐dimensional spectroscopy. Journal of Magnetic Resonance 28: 471–476.

Bodenhausen G and Ruben DJ (1980) Natural abundance nitrogen‐15 NMR by enhanced heteronuclear spectroscopy. Chemical Physics Letters 69: 185–189.

Cheung MS, Maguire ML, Stevens TJ and Broadhurst RW (2010) DANGLE: a Bayesian inferential method for predicting protein backbone dihedral angles and secondary structure. Journal of Magnetic Resonance 202: 223–233.

Demarco A, Llinas M and Wuthrich K (1978) Analysis of the 1H‐NMR spectra of ferrichrome peptides. I. The non‐amide protons. Biopolymers 17: 617–636.

Dickinson WC (1950) Dependence of the F19 nuclear resonance position on chemical compound. Physical Review 77: 736–737.

Dominguez C, Boelens R and Bonvin AMJJ (2003) HADDOCK: a protein–protein docking approach based on biochemical or biophysical information. Journal of the American Chemical Society 125: 1731–1737.

Ernst RR and Anderson WA (1966) Application of Fourier transform spectroscopy to magnetic resonance. Review of Scientific Instruments 37: 93–102.

Fernández C and Wider G (2003) TROSY in NMR studies of the structure and function of large biological macromolecules. Current Opinion in Structural Biology 13: 570–580.

Fiaux J, Bertelsen EB, Horwich AL and Wuthrich K (2002) NMR analysis of a 900K GroEL GroES complex. Nature 418: 207–211.

Griffey RH, Redfield AG, Mcintosh LP, Oas TG and Dahlquist FW (1986) Assignment of proton amide resonances of T4 lysozyme by C‐13 and N‐15 multiple isotopic labeling. Journal of the American Chemical Society 108: 6816–6817.

Grzesiek S, Anglister J and Bax A (1993) Correlation of backbone amide and aliphatic side‐chain resonances in 13C/15N‐enriched proteins by isotropic mixing of 13C magnetization. Journal of Magnetic Resonance B101: 114–119.

Grzesiek S, Cordier F, Jaravine V and Barfield M (2004) Insights into biomolecular hydrogen bonds from hydrogen bond scalar couplings. Progress in NMR Spectroscopy 45: 275–300.

Güntert P (1998) Structure calculation of biological macromolecules from NMR data. Quarterly Reviews of Biophysics 31: 145–237.

Guntert P and Buchner L (2015) Combined automated NOE assignment and structure calculation with CYANA. Journal of Biomolecular NMR 62: 453–471.

Gutowski HS and McCall DW (1951) Nuclear magnetic resonance fine structure in liquids. Physical Review 82: 748–749.

Hahn EL (1950) Spin echoes. Physical Review 80: 580–594.

Ikura M, Kay LE and Bax A (1990) A novel approach for sequential assignment of 1H, 13C, and 15N spectra of proteins: heteronuclear triple‐resonance three‐dimensional NMR spectroscopy. Application to calmodulin. Biochemistry 29: 4659–4667.

Johnson BA (2004) Using NMRview to visualize and analyze the NMR spectra of macromolecules. Methods in Molecular Biology 278: 313–352.

Jung Y‐S and Zweckstetter M (2004) Mars − robust automatic backbone assignment of proteins. Journal of Biomolecular NMR 30: 11–23.

Karplus M (1963) Vicinal proton coupling in nuclear magnetic resonance. Journal of the American Chemical Society 85: 2870–2871.

Kerfah R, Plevin MJ, Sounier R, Gans P and Boisbouvier J (2015) Methyl‐specific isotopic labeling: a molecular tool box for solution NMR studies of large proteins. Current Opinion in Structural Biology 32: 113–122.

Kovacs H, Moskau D and Spraul M (2005) Cryogenically cooled probes − a leap in NMR technology. Progress in NMR Spectroscopy 46: 131–155.

Kumar A, Ernst RR and Wuthrich K (1980) A two‐dimensional nuclear Overhauser enhancement (2D NOE) experiment for the elucidation of complete proton–proton cross‐relaxation networks in biological macromolecules. Biochemical Biophysical Research Communications 95: 1–6.

Laskowski RA, Rullmannn JA, MacArthur MW, Kaptein R and Thornton JM (1996) AQUA and PROCHECK‐NMR: programs for checking the quality of protein structures solved by NMR. Journal of Biomolecular NMR 8: 477–486.

Lee W, Tonelli M and Markley JL (2015) NMRFAM‐SPARKY: enhanced software for biomolecular NMR spectroscopy. Bioinformatics 31: 1325–1327.

Linge JP and Nilges M (1999) Influence of non‐bonded parameters on the quality of NMR structures: a new force field for NMR structure calculation. Journal of Biomolecular NMR 13: 51–59.

Linge JP, Habeck M, Rieping W and Nilges M (2004) Correction of spin diffusion during iterative automated NOE assignment. Journal of Magnetic Resonance 167: 334–342.

Lipsitz RS and Tjandra N (2004) Residual dipolar couplings in NMR structure analysis. Annual Review of Biophysics and Biomolecular Structure 33: 387–413.

Lohr F, Schmidt JM, Maurer S and Ruterjans H (2001) Improved measurement of 3J(Hαi, Ni+1) coupling constants in H2O dissolved proteins. Journal of Magnetic Resonance 153: 75–82.

Mcintosh LP, Muchmore DC, Dahlquist FW, Griffey RH and Redfield AG (1987) Proton NMR studies of bacteriophage T4 lysozyme aided by N‐15 and C‐13 isotopic labeling – a strategy for structural and dynamic investigations of larger proteins. Biophysical Journal 51: A557.

Muchmore DC, McIntosh LP, Russell CB, Anderson DE and Dahlquist FW (1989) Expression and nitrogen‐15 labeling of proteins for proton and nitrogen‐15 nuclear magnetic resonance. Methods in Enzymology 177: 44–73.

Nagayama K, Wuthrich K, Bachmann P and Ernst RR (1977) Two‐dimensional J‐resolved 1H NMR spectroscopy for studies of biological macromolecules. Biochemical and Biophysical Research Communications 78: 99–105.

Oschkinat H, Griesinger C, Kraulis PJ, et al. (1988) Three‐dimensional NMR spectroscopy of a protein in solution. Nature 332: 374–376.

Overhauser AW (1953) Polarization of nuclei in metals. Physical Review 92: 411–415.

Prestegard JH, Bougault CM and Kishore AI (2004) Residual dipolar couplings in structure determination of biomolecules. Chemical Reviews 104: 3519–3540.

Proctor WG and Yu FC (1950) The dependence of a nuclear magnetic resonance frequency upon chemical compound. Physical Review 77: 717.

Purcell EM, Torrey HC and Pound RV (1946) Resonance absorption by nuclear magnetic moments in a solid. Physical Review 69: 37–38.

Rabi II (1937) Space quantization in a gyrating magnetic field. Physical Review 51: 652–654.

Salzmann M, Pervushin K, Wider G, Senn H and Wüthrich K (1998) TROSY in triple‐resonance experiments: new perspectives for sequential NMR assignment of large proteins. Proceedings of the National Academy of Sciences of the United States of America 95: 13585–13590.

Sattler M and Fesik SW (1996) Use of deuterium labeling in NMR: overcoming a sizeable problem. Structure 4: 1245–1249.

Sattler M, Schleucher J and Griesinger C (1999) Heteronuclear multidimensional NMR experiments for the structure determination of proteins in solution employing pulsed field gradients. Progress in NMR Spectroscopy 34: 93–158.

Saunders M, Wishnia A and Kirkwood JG (1957) The nuclear magnetic resonance spectrm of ribonuclease. Journal of the American Chemical Society 79: 3289–3290.

Schmidt JM (2007) Asymmetric Karplus curves for the protein side‐chain 3J couplings. Journal of Biomolecular NMR 37: 287–301.

Shekhtman A, Ghose R, Goger M and Cowburn D (2002) NMR structure determination and investigation using a reduced proton (REDPRO) labelling strategy for proteins. FEBS Letters 524: 177–182.

Shen Y, Delaglio F, Cornilescu G and Bax A (2009) TALOS+: a hybrid method for predicting protein backbone torsion angles from NMR chemical shifts. Journal of Biomolecular NMR 44: 213–223.

Staunton D, Schlinkert R, Zanetti G, Colebrook SA and Campbell ID (2006) Cell‐free expression and selective isotope labelling in protein NMR. Magnetic Resonance in Chemistry 44: S2–S9.

Takahashi H and Shimada I (2010) Production of isotopically labeled heterologous proteins in non‐E. coli prokaryotic and eukaryotic cells. Journal of Biomolecular NMR 46: 3–10.

Tjandra N and Bax A (1997) Direct measurement of distances and angles in biomolecules by NMR in a dilute liquid crystalline medium. Science 278: 1111–1114.

Tolman JR, Flanagan JM, Kennedy MA and Prestegard JH (1995) Nuclear magnetic dipole interactions in field‐oriented proteins: information for structure determination in solution. Proceedings of the National Academy of Sciences of the United States of America 92: 9279–9283.

Tugarinov V and Kay LE (2005) Methyl groups as probes of structure and dynamics in NMR studies of high‐molecular‐weight proteins. ChemBioChem 6: 1567–1577.

Tzakos AG, Grace CRR, Lukavsky PJ and Riek R (2006) NMR techniques for very large proteins and RNAs in solution. Annual Review of Biophysics and Biomolecular Structure 35: 319–342.

Velyvis A and Kay LE (2013) Measurement of Active site ionization equilibria in the 670 kDa proteasome core particle using methyl‐TROSY NMR. Journal of the American Chemical Society 135: 9259–9262.

Vogeli B, Ying J, Grishaev A and Bax A (2007) Limits on variations in protein backbone dynamics from precise measurements of scalar couplings. Journal of the American Chemical Society 129: 9377–9385.

Vranken WF (2014) NMR structure validation in relation to dynamics and structure determination. Progress in NMR Spectroscopy 82: 27–38.

Vriend G (1990) WHAT IF: a molecular modeling and drug design program. Journal of Molecular Graphics 8: 52–56, 29.

Williamson MP, Havel TF and Wuthrich K (1985) Solution conformation of proteinase inhibitor IIA from bull seminal plasma by 1H nuclear magnetic resonance and distance geometry. Journal of Molecular Biology 182: 295–315.

Wishart DS (2011) Interpreting protein chemical shift data. Progress in NMR Spectroscopy 58: 62–87.

Wuthrich K, Wider G, Wagner G and Braun W (1982) Sequential resonance assignments as a basis for determination of spatial protein structures by high resolution proton nuclear magnetic resonance. Journal of Molecular Biology 155: 347–366.

Zhang H and van Ingen H (2016) Isotope‐labeling strategies for solution NMR studies of macromolecular assemblies. Current Opinion in Structural Biology 38: 75–82.

Zuiderweg ERP (2002) Mapping protein–protein interactions in solution by NMR spectroscopy. Biochemistry 41: 1–7.

Further Reading

Cavanagh J, Fairbrother WJ, Palmer AG, Rance M and Skelton JJ (2007) Protein NMR Spectroscopy: Principles and Practice, 2nd edn. New York, NY: Academic Press.

Ernst RR, Bodenhausen G and Wokaun A (1987) Principles of Magnetic Resonance in One and Two Dimensions. Oxford, UK: Clarendon Press.

Ernst RR (1992) Nobel lecture. Nuclear magnetic resonance Fourier transform spectroscopy. Bioscience Reports 12: 143–187.

Ramsey NF (1999) Early history of magnetic resonance. Physics in Perspective 1: 123–135.

Wüthrich K (1986) NMR of Proteins and Nucleic Acids. New York, NY: Wiley Interscience.

Wüthrich K (2003) Nobel Lecture: NMR studies of structure and function of biological macromolecules. Bioscience Reports 23: 119–168.

Nature Milestones – Spin: http://www.nature.com/milestones/milespin/index.html

Related Sub-Topics:

Protein Structure | Proteins: Structure, Function, Metabolism | Nucleic Acids: Structure, Function, Metabolism | Biochemical and Biophysical Techniques | Nuclear Magnetic Resonance | Nucleic Acid Structure
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Article Title: Protein Structure and Interactions from Nuclear Magnetic Resonance Spectroscopy
 
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Ghose, Ranajeet(Jan 2017) Protein Structure and Interactions from Nuclear Magnetic Resonance Spectroscopy. In: eLS. John Wiley & Sons Ltd, Chichester. http://www.els.net [doi: 10.1002/9780470015902.a0003100.pub3]