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Structure and Dynamic Properties of Membrane Proteins using NMR

Heike I. Rösner, Birthe B. Kragelund

10.1002/cphy.c110036

Source: Volume 2, Issue 2, April 2012

Published online: April 2012

Full Article on Wiley Online Library



Abstract

Integral membrane proteins are one of the most challenging groups of macromolecules despite their apparent conformational simplicity. They manage and drive transport, circulate information, and participate in cellular movements via interactions with other proteins and through intricate conformational changes. Their structural and functional decoding is challenging and has imposed demanding experimental development. Solution nuclear magnetic resonance (NMR) spectroscopy is one of the techniques providing the capacity to make a significant difference in the deciphering of the membrane protein structure‐function paradigm. The method has evolved dramatically during the last decade resulting in a plethora of new experiments leading to a significant increase in the scientific repertoire for studying membrane proteins. Besides solving the three‐dimensional structures using state‐of‐the‐art approaches, a large variety of developments of well‐established techniques are available providing insight into membrane protein flexibility, dynamics, and interactions. Inspired by the speed of development in the application of new strategies, by invention of methods to measure solvent accessibility and describe low‐populated states, this review seeks to introduce the vast possibilities solution NMR can offer to the study of membrane protein structure‐function analyses with special focus on applicability. © 2012 American Physiological Society. Compr Physiol 2:1491‐1539, 2012.

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Figure 1. Figure 1.

Structures of 18 selected membrane protein solved by solution NMR spectroscopy. Since 2001, only around 20 structures of integral membrane proteins have been solved in solution and are publicly available. The figure has been divided into two parts, β‐barrel structures in the upper part and α‐helical structures in the lower part. The structures are (A) OmpX, PDB code 1Q9F 78, (B) OmpA, PDB code 2GEA 7, (C) PagP, PDB code 1MM4 111, (D) OmpG, PDB code 2JQY 75, (E) human VDAC‐1, PDB code 2JK4 102, (F) KpOmpA, PDB code 2K0L 119,205, (G) potassium channel KcsA, PDB code 2A9H 10, (H) diacylglycerol kinase, DAGK, PDB code 2KDC 259, (I) phospholamban, PDB code 1ZLL 192, (J) influenza A M2 proton channel, PDB code 2RLF 219, (K) influenza B M2 proton channel, PDB code 2KIX 267, (L) DsbB, PDB code 2K73 289, (M) sensory rhodopsin II, PDB code 2KSY 86, (N) n‐acetylcholine receptor, PDB code 2KSR 28, (O) KCNE1, PDB code 2K21 123, (P) FXYD1, PDB code 2JO1 246, (Q) Rv1761c, PDB code 2K3M 193, and (R) subunit c of the F0F1 ATPase synthase, PDB code 1A91 80. Images of structures generated in PyMol (DeLano Scientific).
Figure 2. Figure 2.

Choice of lipids for NMR characterization. At the top, a schematic illustration of membrane protein extraction and reconstitution into detergents suitable for structural analysis is shown. The detergent needs to preserve the native fold of the internal membrane part. However, some detergents will also affect external parts and even denature those, which is detrimental for structure analysis. Later, an overview of various commonly used detergents applied in solution nuclear magnetic resonance (NMR) studies of IMPs is provided. The chemical structures are grouped according to their nature into organic solvents, nonionic, ionic, zwitterionic, and sterol‐based detergents. For each chemical structure, the critical micelle concentration (cmc) and the aggregation number (AG number) are listed, which are taken either from 215 or from the suppliers.
Figure 3. Figure 3.

The NMR assignment process. Typically by selecting different types of NMR spectra, one selects intra‐residue signals versus inter‐residue signals in a 3D‐spectrum. By a combination of such spectra, which often is nine or more 3D spectra, and together with the knowledge of the sequence of the protein, one performs the job of linking the chemical shifts to distinct nuclei. This process is called assignment. In the assignment process, the NMR signals are linked to the correct nuclei in the protein. (A) A 1D1H‐NMR spectrum of a protein with the chemical shifts of the NMR signals on the x‐axis, approximately going from −1 ppm to 11 ppm. The large overlap between signals demands for extraction into more dimensions. (B) A 2D15N,1H‐HSQC spectrum of a protein with NMR signals from all protons covalently bound to a nitrogen. Here the overlap is minimized dramatically. (C) A 3D HNCO‐spectrum of a protein showing NMR signals from amide protons covalently bound to nitrogen, which again is covalently bound to a carbonyl atom. This type of spectrum requires stable isotope labeling with 15N and 13C. (D) A schematic description of the assignment process. From knowledge of the chemical structure of the peptide chain and the sequence of the protein (E), one uses a set of six to ten 3D heteronuclear NMR spectra to assign the individual NMR signals to the individual atoms of the protein. (F) A schematic representation of the sequential walk, where signals from six different strips of a theoretical HNCACB 3D spectrum are shown. One strip represents carbon signals correlating to a single N,HN. In each strip the N,HN correlates to four carbons, the Cα and Cβ from its own residue and the Cα and Cβ from the preceding residue. This directly links the strips (see arrows). The challenge in assignment is to identify which strip, i.e. which residue is a neighbor to which. (G) An example of the magnetization transfer steps exploited to generate two of the triple‐resonance spectra HNCACB (left) and CBCA(CO)NH (right) important for the assignment. The fist spectrum (left) provides both intraresidual and sequential signals and the second spectrum (right) selects only sequential signals. Arrows indicate the route of transfer and the number the order by which magnetization is transferred from one nucleus to the next. The connections are between residue i and the preceding residue i − 1. All carbons are 13C‐labeled and all nitrogens 15N‐labeled.
Figure 4. Figure 4.

HSQC spectra with and without TROSY‐a tool to circumvent the size problem. Comparison of (A) a conventional 2D [1H, 15N]‐HSQC and (B) a 2D [1H, 15N]‐TROSY spectrum. Both spectra were recorded on a 2 mmol/L OmpX solution in DHPC micelles at 30°C at a resonance frequency of 750 Mhz. The superior performance of the TROSY experiment clearly manifests in the better resolution of the individual cross peaks in the TROSY spectrum. (C) Solution nuclear magnetic resonance (NMR) structure of OmpX in DHPC micelles [20, PDB‐code 1Q9F]. The TROSY effect is largest at very strong magnetic fields. Parts (A) and (B) are modified, with permission, from reference 77. Copyright 2001 National Academy of Sciences USA.
Figure 5. Figure 5.

NMR and the size problem. (A) Molecules tumble freely in solution. However, larger proteins take longer time to tumble around their own axis than small molecules. Left: illustration of a large molecule with slow tumbling, and right: a small molecule with fast tumbling. (B) A nuclear magnetic resonance (NMR) signal is recorded after the application of a radiofrequency pulse that perturbs the equilibrium distribution of spins. One records the return to equilibrium that is an exponential process and is called the free‐induction decay, FID (in red waves). To obtain the NMR spectrum, the FID undergoes a Fourier transformation (FT)—a mathematical operation. For large molecules tumbling more slowly (left), the corresponding NMR signal disappears (relaxes) quickly and the intensity of the signal is dampened, whereas for small molecules (right), the NMR signal is more long‐lived, giving rise to more intense and sharper signals.
Figure 6. Figure 6.

Typical NMR‐derived structural restraints. A number of different conformational restraints can be derived from different types of nuclear magnetic resonance (NMR) experiments. Chemical shifts‐derived dihedral angles restraints: when chemical shifts are assigned to the nuclei, the values are compared to those of databases that link chemical shifts to structures. Hereby one obtains the likelihood for the backbone dihedral angles φ and ψ to be in specific conformations, that is, occupying specific regions of the Ramachandran plot. The dihedral angles are then included together with an allowed angle deviation as input to the structure calculation. NOE‐derived distance restraints: through cross‐relaxation through space, one obtains a distance dependence of the intensity of the NMR signal that declines with the six‐root of the distance. This is termed the NOE effect. This means that only short‐range distances (r), can be detected, typically up to 5 Å. Thus, the intensity of the NOE signal determines the range of the distance restraint, where a strong peak typically is interpreted as a distance being closer than 2.5 Å and weak peak as a distance closer than 5 Å. RDC‐derived global restraints: residual dipolar couplings (RDCs) are global restraints that relate the bond vector to the laboratory frame. From a change in the splitting of the NMR signal, either to larger or smaller splitting, the angles relating the N‐H vector to the laboratory frame can be determined for each vector of the molecule. PRE‐derived distance restraints: paramagnetic relaxation enhancements (PREs) are long‐range distance restraints. The presence of a paramagnetic center enhances the relaxation process and thus lowers the intensity of the NMR signals in its surroundings. For the three cases described here, atom a is closest to the paramagnetic center and thus its NMR signal will be most affected, whereas atom c is too far away to be affected and b is in between.
Figure 7. Figure 7.

Structure calculation scheme. Flowchart describing a possible procedure for protein structure determination by solution nuclear magnetic resonance (NMR) spectroscopy. After protein production and purification a large set of triple resonance, heteronuclear three‐dimensional (3D) NMR spectra are recorded. The chemical shifts are assigned and restraints are derived from chemical shifts, NOEs, PREs, RDCs, etc. Together with information of the chemical structure and sequence, these restraints are fed into a computer algorithm that iteratively calculates and refines the structure and finally yields an ensemble of structures such as shown here for OmpX, fulfilling the experimental restraints. The red arrow indicated the iterative cycle of refinements, which often result in numerous cycles.
Figure 8. Figure 8.

Paramagnetic relaxation enhancement (PRE) and its application in membrane protein structure characterization. PREs can be used to measure long‐range distance restraints in proteins as illustrated in (A) or determine solvent/detergent exposure as illustrated in (B). The 2D [1H, 15N]—HSQC spectrum of Rv1761c in the presence of a (A) reduced and an (B) oxidized MTSL‐label in position F30C. The circles in (B) highlight the residues whose signals are missing in the oxidized spectrum. A more quantitative illustration showing the intensity ratio of reduced (para) and oxidized (dia) cross peaks (Ired/Iox) is shown in (C). An asterisk marks the position of the spin label. From this pattern, additional structural restraints could be deduced. The inset shows the solution structure of Rv1761c with a schematic representation of the influence range of the spin label. (E, F) Schematic illustration of how specifically spin‐labeled detergents can be used to probe the insertion depth in the membrane. (E) Structure 1: 5‐DSA, Structure 2: 16‐DSA. Depending on the position of the spin label within the chemical structure of the detergent, different zones within the transmembrane region will be affected as highlighted by the red rings in (F). Thus, nuclear magnetic resonance (NMR) signals from residues occupying the upper layer are influenced more by 5‐DSA than by 16‐DSA and vice versa.

Parts (A)‐(C) reproduced, with permission, from reference 193. Copyright 2009 John Wiley and Sons.
Figure 9. Figure 9.

Residual dipolar coupling (RDC) and its application in membrane protein structure characterization. (A) Under isotropic conditions the protein is free to tumble in solution and the dipolar coupling is averaged to 0. In this case the splitting in the nuclear magnetic resonance (NMR) signal corresponds to the J‐coupling. (B) In an anisotropic scenario, where submolar fractions of a suitable alignment media has been added to the NMR sample, the splitting of the two peaks changes as a function of the sum of the J‐coupling and the residual dipolar coupling D. (C) With increased alignment the residual dipolar coupling increases and since the sign of the dipolar coupling may be both positive and negative, this changes the splitting in the NMR signal correspondingly. The numbers above the NMR signals are the 1JN‐HN in (A) and the sum of 1JN‐HN and 1DN‐HN in both (B) and (C). The numbers in parentheses are thus (D). All numbers are given in Hz. (D) Refinement of the OmpA structure using RDCs. By using RDC derived restraints, the structure of OmpA improved from a precision of 1.82 ± 0.22 Å, accuracy 2.94 ± 0.18 Å to a precision of 0.90 ± 0.20 Å, accuracy 1.92± 0.15 Å in the absence of additional hydrogen bond restraints and from precision of 1.10 ± 0.15 Å, accuracy 1.66 ± 0.11 Å to a precision of 0.48 ± 0.05 Å, and accuracy 1.02 ± 0.052 Å in the presence of additional hydrogen‐bond restraints. The crystal structure of OmpA is underlying in green, TM regions are shown in blue, and periplasmic loops in red. (E) FXYD1 has a TM helix and an EM helix. Using RDC‐derived restraints, four possible orientations of the EM relative to the TM helix were possible. With the aid of paramagnetic ions probing for solvent exposure, orientation Nr. 1 could be selected. Here, the gradient in red represents the reduction of the signal in the presence of paramagnetic Mn2+ ions in solution and the gray bare the membrane.

Parts (D) reprinted, with permission, from reference 51 and (E) from reference 246. Copyright 2006 and 2007 American Chemical Society.
Figure 10. Figure 10.

Schematic illustration of conformational events in membrane proteins. The motions that change a protein's conformation occur with time‐constants from ps to several minutes, indicated by the red arrow. The fastest motions, vibrations and librations around covalent bonds, result in atoms moving only a fraction of an Ångström. Ligand binding may involve only subtle motions like rearrangement of amino‐acid residues in the binding site or larger movements over several Ångströms‐like domain reorientations. Here, the NMR structure of the extracelluar domain of the 7 TM human corticotrophin‐releasing factor receptor 1 (in blue) complexed with a high affinity agonist in red (PDB code 2L27) 93. Similar movements may be observed for allosteric structural changes where binding of a ligand at one site of the protein changes the structure in another part. This is shown here for phospholamban, where assembly to the pentamer changes the helix curvature (PDB code 2KB7 and 2K0L) 192,248. Lastly, protein folding involves large movements of the whole protein chain as illustrated hypothetical here for the TM domain of the N‐actylcholine receptor B2‐subunit (PDB code 2KSR) 28. Images of structures generated in PyMol (DeLano Scientific).
Figure 11. Figure 11.

Fast time scale dynamics in protein measured by nuclear magnetic resonance (NMR) spectroscopy. (A) The relaxation times T1 (spin‐lattice relaxation time) and T2 (spin‐spin relaxation time) are among other things strongly dependent on the rotational correlation time, τc, of the molecule. Thus for large molecules, the slower it tumbles in solution, the more dominant the influence of the relaxation on the NMR signal becomes. (B) The order parameter S2 is derived from the various different relaxation parameters and yields a measure for the dependent local mobility of the NMR probe. One qualitative way to describe the order parameter is to describe the motion the bond vector as a cone with an angular dependence θ (C). The smaller this cone, the larger the generalized order parameter S2. Thus for fully rigid structures, S2 will be 1, and for completely random structures S2 will be 0. Often S2 is decomposed into two separate contributions, S2f, which is the fast component representing the ps dynamics and S2s the slow component representing the ns dynamics. Membranes influence IMP dynamics. The water‐soluble analogue WSK3 to the potassium channel KcsA was analyzed by [1H, 15N]—relaxation parameters at 14.1 T(black) and 16.5 T (red). The experimental data are shown in panel (D). These were fitted to the extended model free approach yielding Sf2, SS2, and τs shown in (E). The water‐soluble WSK3 shows dramatically increased dynamics in the slow time regime as represented by SS2 when compared to the original values for the dynamics of KcsA. Especially the pore helix appears to be affected as highlighted in red in (F) (top: KcsA, bottom: WSK3).

Figures (D) and (E) reproduced, with permission, from reference 159. Copyright 2008 National Academy of Sciences USA.
Figure 12. Figure 12.

The influence of chemical exchange on the nuclear magnetic resonance (NMR) signal. Slow chemical exchange results in two distinct signals to be recorded (A top). Chemical exchange on the millisecond (ms) to microsecond (μs) timescale will result in line broadening of the NMR signal which in the extreme case results in line broadening beyond detection (A middle). For chemical exchange in the fast regime, an averaged signal will be recorded (A bottom). In the last case, the position of the signal will depend on the population ratio of the interconverting species and the interconversion rate as illustrated in (B) (C) Chemical exchange also contributes to an apparent faster loss of the signal (R2 + Rex) than can be attributed to the relaxation process described by R2 alone. (D) This loss can be partially recovered by applying a train of refocusing pulses, a so‐called refocusing CPMG pulse sequence in the preparation time. (E) The combined relaxation rate as a function of the frequency of the CPMG sequence. The CPMG will thus remove the contribution from Rex to R2and thereby yield information on the underlying chemical exchange rate Rex. (F) PagP exists in a relaxed (R) and a tensed (T) state, and shows two different NMR signals at 25°C. These two sets of peaks show distinct dynamic differences on the intermediate timescale as highlighted for the residues Y38 and Q139 in (G). (H) Chemical shift differences between the R and the T states color coded onto the cartoon structure of PagP (PDB code 1MM4). (I) The ensemble of the 20 lowest energy structures of PagP highlighting the flexibility of the EM parts.

Parts (F)‐(H) reproduced, with permission, from reference 110. Copyright 2004 National Academy of Sciences USA
Figure 13. Figure 13.

Amide hydrogen to deuterium exchange measured by solution nuclear magnetic resonance (NMR) spectroscopy. (A) Exchange of amides in a membrane protein requires access to bulk solvent, either directly or through conformational changes. An amide proton exchanges with an intrinsic rate constant dependent on the chemical structure of the peptide chain, kint, and with an observed exchange rate constant, kobs, which depends on the structural environment. (B) Since the deuteron has different NMR properties than the proton, it will not be detected in a proton NMR spectrum and as the amide at a certain position is exchanged with a deuteron, the signal gradually disappears following an exponential decay (D). In the lower square (E), two different amides with different rates of exchange are shown, one with fast (red) and one with slower (black) exchange. Each data point represents the intensity of an NMR peak in an HSQC spectrum. (C) A protection factor P can be described as the ratio of the intrinsic to the observed rate constant. The more protected the amide, the larger the protection factor. A log scaling is typically used to illustrate P. For membrane proteins an equilibrium factor is often used, ɛeq, which is the intensity ratio of the NMR signal in H2O to that of the NMR signal in a given amount of D2O. Here, the larger the equilibrium factor, the less protected the amide is. (D) Equilibrium factors used to describe the water accessibility of OmpX. The equilibrium exchange factor ɛeq represents the ratio of peak volumes for OmpX/APol complexes incubated in 100 versus 5% D2O. The ɛeq is normalized relative to the ratio observed for the most protected residue. Here diversity in exchange factors was seen within the barrel, with some strands in the middle of the membrane being most protected while other strands show increased exchange (lower) in the membrane center. This suggesting that dynamics in the barrel is important for the function of OmpX 39.

Part (F) reproduced, with permission, from reference 39. Copyright 2009 Springer.
Figure 14. Figure 14.

Single‐pass IMP structures solved by solution nuclear magnetic resonance (NMR) spectroscopy. (A) Glycoprotein E1 from hepatitis C (PDB code 1EMZ) 186; in red is shown three glycine residues, (B) Sarcolipin, a regulator of Ca‐ATPase (PDB code 1JDM) 165; in red is shown the only three polar residues, (C) KCNE1 a modulator of the KCNQ1 potassium channel (PDB code 2K21) 123, (D) The tetrameric M2‐channel from influenza A virus (PDB code 2RLF) 219; shown in red are four tryptophans and in green above four histidines and in green below four aspartic acids, (E) EphA1 (PDB code 1KIL) 23, in red is shown the tandem glycine motif, and in green a leucine zipper motif facing the lipids, (F) EphA2 (PDB code 2K9Y) 24, in red the extensive hydrophobic interface ending with a Phe‐stack, (G) neu/ErbB2 monomer (PDB code 1IIJ) 91, (H) ErbB2 homodimer (PDB code 2JWA) 25; in red the Ser/Thr‐node, in green valines and leucines as well as valines in blue sticks facing the lipids, (I) ErbB1:ErbB2 heterodimer (PDB code 2KS1) 175; in red the GXXXG:Ser/Thr packing, (J) αIIbβ3‐heterodimer (PDB code 2KNC) 281, in red glycines and in green a tryptophan, (K) Glycophorin A (PDB code1AFO) 160, in red the glycines and valines of the glycine zipper motif, (L) BNip3 homodimer (PDB code 2J5D) 26, in red at the top the Phe‐cluster, at bottom the GXXXG‐motif, in green the Ser‐His node, (M) ζζ‐homodimer (PDB code 2HAC) 37, in red the Tyr/Thr network, in green the Asp‐lock, (N) DAP12 homodimer (PDB code 2L34) 38, in red at the top the GXXXG‐motif, in bottom the threonines, in green the Asp‐lock, and (O) DAP12:NGK2C heterotrimer (PDB code 2L35) 38. In red the NGK2C helix, in green the Lys:Asp network. The three monomer structures were solved in TFE or sodium dodecyl sulphate (SDS). Images of structures generated in PyMol (DeLano Scientific).
Figure 15. Figure 15.

Fragments of IMPs can be fair structural representatives. The entire sequence of bacteriorhodopsin was synthesized as short, overlapping peptides, and their structures solved by solution NMR spectroscopy. These structures were compared to the crystal structure and showed surprising similarity throughout, also what concerned turn structures.

The figure is reproduced from the Biophysical Journal 124, with permission, from Elsevier.
Figure 16. Figure 16.

Interacellular juxtamembrane structures of integral membrane proteins (IMPs) solved by solution nuclear magnetic resonance (NMR) spectroscopy. (A) For some intracellular domains of IMPs, an equilibrium between a solvent resident, disordered state and a membrane embedded, helical state is suggested and its importance in receptor regulation presently discussed. (B) Structure of the αIIbβ3‐complex (PDB code 1M8O) 264 and those of the monomeric states of αIIb (PDB code 1DPK) and of β3 (pdb code 1S4X) 263, as well as of an extended version of β3 (PDB code 2KV9) 173. (C) The ICD of the epidermal growth receptor (EGF)‐receptor forms a similar EM structure as β3 (PDB code 1Z9I) 48. (D) Structure of human CD4mut with the TM helix and part of the ICD (PDB code 2KLU) 278. (E) The phospholamban monomer (PDB code 2KB7) 248. (F) The ICD of the T‐cell receptor CD3ɛ with two ITAM motifs (PDB code 2K4F) 280; the tyrosine and leucines are shown in sticks. (G) Mec4 (PDB code 2K2B). JM part of viruses and bacteriophages proteins (H) HIV‐1 virus protein U (VpU) with tertiary contact between EM helices (PDB code 2K7Y) 276, (I) Hepatitis C glycoprotein E1, HepC E1 (PDB code 2KNU) 234, (J) fd coat protein from filamentous bacteriophages (pdb code 1FDM) 4, and (K) tryptophan‐rich domain of HIV glycoprotein gp41 (PDB code 1JAV) 218. Images of structures generated in PyMol (DeLano Scientific).
Figure 17. Figure 17.

Structure‐activity relationship (SAR) by nuclear magnetic resonance (NMR). (A) Cartoon describing the pipeline of novel lead candidate generation. Small probes are used to screen the binding site and are subsequently combined to form the new lead compound. (B) A summary of the approach to find novel p‐arylthio cinnamides as lead candidates for the leukocyte function associated antigen 1 (LFA‐1) 152. The different chemical compounds identified in the various screening steps are drawn as chemical structures under the respective screen steps.
Figure 18. Figure 18.

Nuclear magnetic resonance (NMR) shift mapping—a tool to locate binding sites and interactions. Chemical shift perturbations of CR56 and RAPd1 by complex formation. (A) HSQC spectra of free (black peaks) and CR56‐bound (red) RAPd1. (B) Combined chemical shift changes ((|ΔδH| + 1/4|ΔδN|)0.5) are shown as bars versus residue number for 15N‐CR56 upon binding of unlabeled RAPd1. Proline residues (+) and nonassigned residues in the bound form (x) are indicated. The CR5 and CR6 domain boundaries are indicated with bars above the plots. (C) Surface plots showing in red those residues with significantly perturbed chemical shifts upon complex formation in CR56 (Ala944, Cys948, Trp953, Thr954, Asp956, Asp966, Ser968, Ser970, Ala977, Thr975, Phe977, Thr980, Gly987, Asp997, Asn998, and Gly1003). (D) Sections of 15N‐1H HSQC spectra showing the peaks from RAPd1 residue Glu40 during the titration of 15N‐labeled RAPd1 with increasing amounts of unlabeled CR56. Figure modified, with permission, from reference 120 and with permission from Elsevier.
Figure 19. Figure 19.

Ligand screening methods. A summary of the various NMR methods used to screen for ligands with moderately binding affinity, highlighting subtle differences in the modes of action. Paramagnetic relaxation enhancement (PRE): a label is attached to the macromolecule close to the ligand binding site and will modify the relaxation properties of nearby residues, including atoms of a weakly interacting ligand. Diffusion‐ordered spectroscopy (DOSY): the interaction of the ligand with the protein will result into slower diffusion properties of the ligand than expected from its actual size, resulting in modification of the NMR parameters of the ligand. Saturation transfer difference (STD) and trNOE: saturation will be transferred from the macromolecule to the ligand. The efficiency of this transfer will depend on the actual orientation of the ligand when bound to the protein. Interligand nuclear Overhauser effect (ILOE): the interligand NOE makes use of the strong negative ligand‐ligand NOE. The NOEs can be used to identify those different ligands that bind simultaneously and can help in the design of novel lead candidates based on the initial two weak binders and their orientation relative to each other.


Figure 1.

Structures of 18 selected membrane protein solved by solution NMR spectroscopy. Since 2001, only around 20 structures of integral membrane proteins have been solved in solution and are publicly available. The figure has been divided into two parts, β‐barrel structures in the upper part and α‐helical structures in the lower part. The structures are (A) OmpX, PDB code 1Q9F 78, (B) OmpA, PDB code 2GEA 7, (C) PagP, PDB code 1MM4 111, (D) OmpG, PDB code 2JQY 75, (E) human VDAC‐1, PDB code 2JK4 102, (F) KpOmpA, PDB code 2K0L 119,205, (G) potassium channel KcsA, PDB code 2A9H 10, (H) diacylglycerol kinase, DAGK, PDB code 2KDC 259, (I) phospholamban, PDB code 1ZLL 192, (J) influenza A M2 proton channel, PDB code 2RLF 219, (K) influenza B M2 proton channel, PDB code 2KIX 267, (L) DsbB, PDB code 2K73 289, (M) sensory rhodopsin II, PDB code 2KSY 86, (N) n‐acetylcholine receptor, PDB code 2KSR 28, (O) KCNE1, PDB code 2K21 123, (P) FXYD1, PDB code 2JO1 246, (Q) Rv1761c, PDB code 2K3M 193, and (R) subunit c of the F0F1 ATPase synthase, PDB code 1A91 80. Images of structures generated in PyMol (DeLano Scientific).



Figure 2.

Choice of lipids for NMR characterization. At the top, a schematic illustration of membrane protein extraction and reconstitution into detergents suitable for structural analysis is shown. The detergent needs to preserve the native fold of the internal membrane part. However, some detergents will also affect external parts and even denature those, which is detrimental for structure analysis. Later, an overview of various commonly used detergents applied in solution nuclear magnetic resonance (NMR) studies of IMPs is provided. The chemical structures are grouped according to their nature into organic solvents, nonionic, ionic, zwitterionic, and sterol‐based detergents. For each chemical structure, the critical micelle concentration (cmc) and the aggregation number (AG number) are listed, which are taken either from 215 or from the suppliers.



Figure 3.

The NMR assignment process. Typically by selecting different types of NMR spectra, one selects intra‐residue signals versus inter‐residue signals in a 3D‐spectrum. By a combination of such spectra, which often is nine or more 3D spectra, and together with the knowledge of the sequence of the protein, one performs the job of linking the chemical shifts to distinct nuclei. This process is called assignment. In the assignment process, the NMR signals are linked to the correct nuclei in the protein. (A) A 1D1H‐NMR spectrum of a protein with the chemical shifts of the NMR signals on the x‐axis, approximately going from −1 ppm to 11 ppm. The large overlap between signals demands for extraction into more dimensions. (B) A 2D15N,1H‐HSQC spectrum of a protein with NMR signals from all protons covalently bound to a nitrogen. Here the overlap is minimized dramatically. (C) A 3D HNCO‐spectrum of a protein showing NMR signals from amide protons covalently bound to nitrogen, which again is covalently bound to a carbonyl atom. This type of spectrum requires stable isotope labeling with 15N and 13C. (D) A schematic description of the assignment process. From knowledge of the chemical structure of the peptide chain and the sequence of the protein (E), one uses a set of six to ten 3D heteronuclear NMR spectra to assign the individual NMR signals to the individual atoms of the protein. (F) A schematic representation of the sequential walk, where signals from six different strips of a theoretical HNCACB 3D spectrum are shown. One strip represents carbon signals correlating to a single N,HN. In each strip the N,HN correlates to four carbons, the Cα and Cβ from its own residue and the Cα and Cβ from the preceding residue. This directly links the strips (see arrows). The challenge in assignment is to identify which strip, i.e. which residue is a neighbor to which. (G) An example of the magnetization transfer steps exploited to generate two of the triple‐resonance spectra HNCACB (left) and CBCA(CO)NH (right) important for the assignment. The fist spectrum (left) provides both intraresidual and sequential signals and the second spectrum (right) selects only sequential signals. Arrows indicate the route of transfer and the number the order by which magnetization is transferred from one nucleus to the next. The connections are between residue i and the preceding residue i − 1. All carbons are 13C‐labeled and all nitrogens 15N‐labeled.



Figure 4.

HSQC spectra with and without TROSY‐a tool to circumvent the size problem. Comparison of (A) a conventional 2D [1H, 15N]‐HSQC and (B) a 2D [1H, 15N]‐TROSY spectrum. Both spectra were recorded on a 2 mmol/L OmpX solution in DHPC micelles at 30°C at a resonance frequency of 750 Mhz. The superior performance of the TROSY experiment clearly manifests in the better resolution of the individual cross peaks in the TROSY spectrum. (C) Solution nuclear magnetic resonance (NMR) structure of OmpX in DHPC micelles [20, PDB‐code 1Q9F]. The TROSY effect is largest at very strong magnetic fields. Parts (A) and (B) are modified, with permission, from reference 77. Copyright 2001 National Academy of Sciences USA.



Figure 5.

NMR and the size problem. (A) Molecules tumble freely in solution. However, larger proteins take longer time to tumble around their own axis than small molecules. Left: illustration of a large molecule with slow tumbling, and right: a small molecule with fast tumbling. (B) A nuclear magnetic resonance (NMR) signal is recorded after the application of a radiofrequency pulse that perturbs the equilibrium distribution of spins. One records the return to equilibrium that is an exponential process and is called the free‐induction decay, FID (in red waves). To obtain the NMR spectrum, the FID undergoes a Fourier transformation (FT)—a mathematical operation. For large molecules tumbling more slowly (left), the corresponding NMR signal disappears (relaxes) quickly and the intensity of the signal is dampened, whereas for small molecules (right), the NMR signal is more long‐lived, giving rise to more intense and sharper signals.



Figure 6.

Typical NMR‐derived structural restraints. A number of different conformational restraints can be derived from different types of nuclear magnetic resonance (NMR) experiments. Chemical shifts‐derived dihedral angles restraints: when chemical shifts are assigned to the nuclei, the values are compared to those of databases that link chemical shifts to structures. Hereby one obtains the likelihood for the backbone dihedral angles φ and ψ to be in specific conformations, that is, occupying specific regions of the Ramachandran plot. The dihedral angles are then included together with an allowed angle deviation as input to the structure calculation. NOE‐derived distance restraints: through cross‐relaxation through space, one obtains a distance dependence of the intensity of the NMR signal that declines with the six‐root of the distance. This is termed the NOE effect. This means that only short‐range distances (r), can be detected, typically up to 5 Å. Thus, the intensity of the NOE signal determines the range of the distance restraint, where a strong peak typically is interpreted as a distance being closer than 2.5 Å and weak peak as a distance closer than 5 Å. RDC‐derived global restraints: residual dipolar couplings (RDCs) are global restraints that relate the bond vector to the laboratory frame. From a change in the splitting of the NMR signal, either to larger or smaller splitting, the angles relating the N‐H vector to the laboratory frame can be determined for each vector of the molecule. PRE‐derived distance restraints: paramagnetic relaxation enhancements (PREs) are long‐range distance restraints. The presence of a paramagnetic center enhances the relaxation process and thus lowers the intensity of the NMR signals in its surroundings. For the three cases described here, atom a is closest to the paramagnetic center and thus its NMR signal will be most affected, whereas atom c is too far away to be affected and b is in between.



Figure 7.

Structure calculation scheme. Flowchart describing a possible procedure for protein structure determination by solution nuclear magnetic resonance (NMR) spectroscopy. After protein production and purification a large set of triple resonance, heteronuclear three‐dimensional (3D) NMR spectra are recorded. The chemical shifts are assigned and restraints are derived from chemical shifts, NOEs, PREs, RDCs, etc. Together with information of the chemical structure and sequence, these restraints are fed into a computer algorithm that iteratively calculates and refines the structure and finally yields an ensemble of structures such as shown here for OmpX, fulfilling the experimental restraints. The red arrow indicated the iterative cycle of refinements, which often result in numerous cycles.



Figure 8.

Paramagnetic relaxation enhancement (PRE) and its application in membrane protein structure characterization. PREs can be used to measure long‐range distance restraints in proteins as illustrated in (A) or determine solvent/detergent exposure as illustrated in (B). The 2D [1H, 15N]—HSQC spectrum of Rv1761c in the presence of a (A) reduced and an (B) oxidized MTSL‐label in position F30C. The circles in (B) highlight the residues whose signals are missing in the oxidized spectrum. A more quantitative illustration showing the intensity ratio of reduced (para) and oxidized (dia) cross peaks (Ired/Iox) is shown in (C). An asterisk marks the position of the spin label. From this pattern, additional structural restraints could be deduced. The inset shows the solution structure of Rv1761c with a schematic representation of the influence range of the spin label. (E, F) Schematic illustration of how specifically spin‐labeled detergents can be used to probe the insertion depth in the membrane. (E) Structure 1: 5‐DSA, Structure 2: 16‐DSA. Depending on the position of the spin label within the chemical structure of the detergent, different zones within the transmembrane region will be affected as highlighted by the red rings in (F). Thus, nuclear magnetic resonance (NMR) signals from residues occupying the upper layer are influenced more by 5‐DSA than by 16‐DSA and vice versa.

Parts (A)‐(C) reproduced, with permission, from reference 193. Copyright 2009 John Wiley and Sons.


Figure 9.

Residual dipolar coupling (RDC) and its application in membrane protein structure characterization. (A) Under isotropic conditions the protein is free to tumble in solution and the dipolar coupling is averaged to 0. In this case the splitting in the nuclear magnetic resonance (NMR) signal corresponds to the J‐coupling. (B) In an anisotropic scenario, where submolar fractions of a suitable alignment media has been added to the NMR sample, the splitting of the two peaks changes as a function of the sum of the J‐coupling and the residual dipolar coupling D. (C) With increased alignment the residual dipolar coupling increases and since the sign of the dipolar coupling may be both positive and negative, this changes the splitting in the NMR signal correspondingly. The numbers above the NMR signals are the 1JN‐HN in (A) and the sum of 1JN‐HN and 1DN‐HN in both (B) and (C). The numbers in parentheses are thus (D). All numbers are given in Hz. (D) Refinement of the OmpA structure using RDCs. By using RDC derived restraints, the structure of OmpA improved from a precision of 1.82 ± 0.22 Å, accuracy 2.94 ± 0.18 Å to a precision of 0.90 ± 0.20 Å, accuracy 1.92± 0.15 Å in the absence of additional hydrogen bond restraints and from precision of 1.10 ± 0.15 Å, accuracy 1.66 ± 0.11 Å to a precision of 0.48 ± 0.05 Å, and accuracy 1.02 ± 0.052 Å in the presence of additional hydrogen‐bond restraints. The crystal structure of OmpA is underlying in green, TM regions are shown in blue, and periplasmic loops in red. (E) FXYD1 has a TM helix and an EM helix. Using RDC‐derived restraints, four possible orientations of the EM relative to the TM helix were possible. With the aid of paramagnetic ions probing for solvent exposure, orientation Nr. 1 could be selected. Here, the gradient in red represents the reduction of the signal in the presence of paramagnetic Mn2+ ions in solution and the gray bare the membrane.

Parts (D) reprinted, with permission, from reference 51 and (E) from reference 246. Copyright 2006 and 2007 American Chemical Society.


Figure 10.

Schematic illustration of conformational events in membrane proteins. The motions that change a protein's conformation occur with time‐constants from ps to several minutes, indicated by the red arrow. The fastest motions, vibrations and librations around covalent bonds, result in atoms moving only a fraction of an Ångström. Ligand binding may involve only subtle motions like rearrangement of amino‐acid residues in the binding site or larger movements over several Ångströms‐like domain reorientations. Here, the NMR structure of the extracelluar domain of the 7 TM human corticotrophin‐releasing factor receptor 1 (in blue) complexed with a high affinity agonist in red (PDB code 2L27) 93. Similar movements may be observed for allosteric structural changes where binding of a ligand at one site of the protein changes the structure in another part. This is shown here for phospholamban, where assembly to the pentamer changes the helix curvature (PDB code 2KB7 and 2K0L) 192,248. Lastly, protein folding involves large movements of the whole protein chain as illustrated hypothetical here for the TM domain of the N‐actylcholine receptor B2‐subunit (PDB code 2KSR) 28. Images of structures generated in PyMol (DeLano Scientific).



Figure 11.

Fast time scale dynamics in protein measured by nuclear magnetic resonance (NMR) spectroscopy. (A) The relaxation times T1 (spin‐lattice relaxation time) and T2 (spin‐spin relaxation time) are among other things strongly dependent on the rotational correlation time, τc, of the molecule. Thus for large molecules, the slower it tumbles in solution, the more dominant the influence of the relaxation on the NMR signal becomes. (B) The order parameter S2 is derived from the various different relaxation parameters and yields a measure for the dependent local mobility of the NMR probe. One qualitative way to describe the order parameter is to describe the motion the bond vector as a cone with an angular dependence θ (C). The smaller this cone, the larger the generalized order parameter S2. Thus for fully rigid structures, S2 will be 1, and for completely random structures S2 will be 0. Often S2 is decomposed into two separate contributions, S2f, which is the fast component representing the ps dynamics and S2s the slow component representing the ns dynamics. Membranes influence IMP dynamics. The water‐soluble analogue WSK3 to the potassium channel KcsA was analyzed by [1H, 15N]—relaxation parameters at 14.1 T(black) and 16.5 T (red). The experimental data are shown in panel (D). These were fitted to the extended model free approach yielding Sf2, SS2, and τs shown in (E). The water‐soluble WSK3 shows dramatically increased dynamics in the slow time regime as represented by SS2 when compared to the original values for the dynamics of KcsA. Especially the pore helix appears to be affected as highlighted in red in (F) (top: KcsA, bottom: WSK3).

Figures (D) and (E) reproduced, with permission, from reference 159. Copyright 2008 National Academy of Sciences USA.


Figure 12.

The influence of chemical exchange on the nuclear magnetic resonance (NMR) signal. Slow chemical exchange results in two distinct signals to be recorded (A top). Chemical exchange on the millisecond (ms) to microsecond (μs) timescale will result in line broadening of the NMR signal which in the extreme case results in line broadening beyond detection (A middle). For chemical exchange in the fast regime, an averaged signal will be recorded (A bottom). In the last case, the position of the signal will depend on the population ratio of the interconverting species and the interconversion rate as illustrated in (B) (C) Chemical exchange also contributes to an apparent faster loss of the signal (R2 + Rex) than can be attributed to the relaxation process described by R2 alone. (D) This loss can be partially recovered by applying a train of refocusing pulses, a so‐called refocusing CPMG pulse sequence in the preparation time. (E) The combined relaxation rate as a function of the frequency of the CPMG sequence. The CPMG will thus remove the contribution from Rex to R2and thereby yield information on the underlying chemical exchange rate Rex. (F) PagP exists in a relaxed (R) and a tensed (T) state, and shows two different NMR signals at 25°C. These two sets of peaks show distinct dynamic differences on the intermediate timescale as highlighted for the residues Y38 and Q139 in (G). (H) Chemical shift differences between the R and the T states color coded onto the cartoon structure of PagP (PDB code 1MM4). (I) The ensemble of the 20 lowest energy structures of PagP highlighting the flexibility of the EM parts.

Parts (F)‐(H) reproduced, with permission, from reference 110. Copyright 2004 National Academy of Sciences USA


Figure 13.

Amide hydrogen to deuterium exchange measured by solution nuclear magnetic resonance (NMR) spectroscopy. (A) Exchange of amides in a membrane protein requires access to bulk solvent, either directly or through conformational changes. An amide proton exchanges with an intrinsic rate constant dependent on the chemical structure of the peptide chain, kint, and with an observed exchange rate constant, kobs, which depends on the structural environment. (B) Since the deuteron has different NMR properties than the proton, it will not be detected in a proton NMR spectrum and as the amide at a certain position is exchanged with a deuteron, the signal gradually disappears following an exponential decay (D). In the lower square (E), two different amides with different rates of exchange are shown, one with fast (red) and one with slower (black) exchange. Each data point represents the intensity of an NMR peak in an HSQC spectrum. (C) A protection factor P can be described as the ratio of the intrinsic to the observed rate constant. The more protected the amide, the larger the protection factor. A log scaling is typically used to illustrate P. For membrane proteins an equilibrium factor is often used, ɛeq, which is the intensity ratio of the NMR signal in H2O to that of the NMR signal in a given amount of D2O. Here, the larger the equilibrium factor, the less protected the amide is. (D) Equilibrium factors used to describe the water accessibility of OmpX. The equilibrium exchange factor ɛeq represents the ratio of peak volumes for OmpX/APol complexes incubated in 100 versus 5% D2O. The ɛeq is normalized relative to the ratio observed for the most protected residue. Here diversity in exchange factors was seen within the barrel, with some strands in the middle of the membrane being most protected while other strands show increased exchange (lower) in the membrane center. This suggesting that dynamics in the barrel is important for the function of OmpX 39.

Part (F) reproduced, with permission, from reference 39. Copyright 2009 Springer.


Figure 14.

Single‐pass IMP structures solved by solution nuclear magnetic resonance (NMR) spectroscopy. (A) Glycoprotein E1 from hepatitis C (PDB code 1EMZ) 186; in red is shown three glycine residues, (B) Sarcolipin, a regulator of Ca‐ATPase (PDB code 1JDM) 165; in red is shown the only three polar residues, (C) KCNE1 a modulator of the KCNQ1 potassium channel (PDB code 2K21) 123, (D) The tetrameric M2‐channel from influenza A virus (PDB code 2RLF) 219; shown in red are four tryptophans and in green above four histidines and in green below four aspartic acids, (E) EphA1 (PDB code 1KIL) 23, in red is shown the tandem glycine motif, and in green a leucine zipper motif facing the lipids, (F) EphA2 (PDB code 2K9Y) 24, in red the extensive hydrophobic interface ending with a Phe‐stack, (G) neu/ErbB2 monomer (PDB code 1IIJ) 91, (H) ErbB2 homodimer (PDB code 2JWA) 25; in red the Ser/Thr‐node, in green valines and leucines as well as valines in blue sticks facing the lipids, (I) ErbB1:ErbB2 heterodimer (PDB code 2KS1) 175; in red the GXXXG:Ser/Thr packing, (J) αIIbβ3‐heterodimer (PDB code 2KNC) 281, in red glycines and in green a tryptophan, (K) Glycophorin A (PDB code1AFO) 160, in red the glycines and valines of the glycine zipper motif, (L) BNip3 homodimer (PDB code 2J5D) 26, in red at the top the Phe‐cluster, at bottom the GXXXG‐motif, in green the Ser‐His node, (M) ζζ‐homodimer (PDB code 2HAC) 37, in red the Tyr/Thr network, in green the Asp‐lock, (N) DAP12 homodimer (PDB code 2L34) 38, in red at the top the GXXXG‐motif, in bottom the threonines, in green the Asp‐lock, and (O) DAP12:NGK2C heterotrimer (PDB code 2L35) 38. In red the NGK2C helix, in green the Lys:Asp network. The three monomer structures were solved in TFE or sodium dodecyl sulphate (SDS). Images of structures generated in PyMol (DeLano Scientific).



Figure 15.

Fragments of IMPs can be fair structural representatives. The entire sequence of bacteriorhodopsin was synthesized as short, overlapping peptides, and their structures solved by solution NMR spectroscopy. These structures were compared to the crystal structure and showed surprising similarity throughout, also what concerned turn structures.

The figure is reproduced from the Biophysical Journal 124, with permission, from Elsevier.


Figure 16.

Interacellular juxtamembrane structures of integral membrane proteins (IMPs) solved by solution nuclear magnetic resonance (NMR) spectroscopy. (A) For some intracellular domains of IMPs, an equilibrium between a solvent resident, disordered state and a membrane embedded, helical state is suggested and its importance in receptor regulation presently discussed. (B) Structure of the αIIbβ3‐complex (PDB code 1M8O) 264 and those of the monomeric states of αIIb (PDB code 1DPK) and of β3 (pdb code 1S4X) 263, as well as of an extended version of β3 (PDB code 2KV9) 173. (C) The ICD of the epidermal growth receptor (EGF)‐receptor forms a similar EM structure as β3 (PDB code 1Z9I) 48. (D) Structure of human CD4mut with the TM helix and part of the ICD (PDB code 2KLU) 278. (E) The phospholamban monomer (PDB code 2KB7) 248. (F) The ICD of the T‐cell receptor CD3ɛ with two ITAM motifs (PDB code 2K4F) 280; the tyrosine and leucines are shown in sticks. (G) Mec4 (PDB code 2K2B). JM part of viruses and bacteriophages proteins (H) HIV‐1 virus protein U (VpU) with tertiary contact between EM helices (PDB code 2K7Y) 276, (I) Hepatitis C glycoprotein E1, HepC E1 (PDB code 2KNU) 234, (J) fd coat protein from filamentous bacteriophages (pdb code 1FDM) 4, and (K) tryptophan‐rich domain of HIV glycoprotein gp41 (PDB code 1JAV) 218. Images of structures generated in PyMol (DeLano Scientific).



Figure 17.

Structure‐activity relationship (SAR) by nuclear magnetic resonance (NMR). (A) Cartoon describing the pipeline of novel lead candidate generation. Small probes are used to screen the binding site and are subsequently combined to form the new lead compound. (B) A summary of the approach to find novel p‐arylthio cinnamides as lead candidates for the leukocyte function associated antigen 1 (LFA‐1) 152. The different chemical compounds identified in the various screening steps are drawn as chemical structures under the respective screen steps.



Figure 18.

Nuclear magnetic resonance (NMR) shift mapping—a tool to locate binding sites and interactions. Chemical shift perturbations of CR56 and RAPd1 by complex formation. (A) HSQC spectra of free (black peaks) and CR56‐bound (red) RAPd1. (B) Combined chemical shift changes ((|ΔδH| + 1/4|ΔδN|)0.5) are shown as bars versus residue number for 15N‐CR56 upon binding of unlabeled RAPd1. Proline residues (+) and nonassigned residues in the bound form (x) are indicated. The CR5 and CR6 domain boundaries are indicated with bars above the plots. (C) Surface plots showing in red those residues with significantly perturbed chemical shifts upon complex formation in CR56 (Ala944, Cys948, Trp953, Thr954, Asp956, Asp966, Ser968, Ser970, Ala977, Thr975, Phe977, Thr980, Gly987, Asp997, Asn998, and Gly1003). (D) Sections of 15N‐1H HSQC spectra showing the peaks from RAPd1 residue Glu40 during the titration of 15N‐labeled RAPd1 with increasing amounts of unlabeled CR56. Figure modified, with permission, from reference 120 and with permission from Elsevier.



Figure 19.

Ligand screening methods. A summary of the various NMR methods used to screen for ligands with moderately binding affinity, highlighting subtle differences in the modes of action. Paramagnetic relaxation enhancement (PRE): a label is attached to the macromolecule close to the ligand binding site and will modify the relaxation properties of nearby residues, including atoms of a weakly interacting ligand. Diffusion‐ordered spectroscopy (DOSY): the interaction of the ligand with the protein will result into slower diffusion properties of the ligand than expected from its actual size, resulting in modification of the NMR parameters of the ligand. Saturation transfer difference (STD) and trNOE: saturation will be transferred from the macromolecule to the ligand. The efficiency of this transfer will depend on the actual orientation of the ligand when bound to the protein. Interligand nuclear Overhauser effect (ILOE): the interligand NOE makes use of the strong negative ligand‐ligand NOE. The NOEs can be used to identify those different ligands that bind simultaneously and can help in the design of novel lead candidates based on the initial two weak binders and their orientation relative to each other.

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Article Title: Structure and Dynamic Properties of Membrane Proteins using NMR
 

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Heike I. Rösner, Birthe B. Kragelund. Structure and Dynamic Properties of Membrane Proteins using NMR. Compr Physiol 2012, 2: 1491-1539. doi: 10.1002/cphy.c110036