Recent advances in magic angle spinning solid state NMR of membrane proteins.

Progress in Nuclear Magnetic Resonance Spectroscopy 82 (2014) 1–26

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Progress in Nuclear Magnetic Resonance Spectroscopy journal homepage: www.elsevier.com/locate/pnmrs

Recent advances in magic angle spinning solid state NMR of membrane proteins Shenlin Wang a,b, Vladimir Ladizhansky c,d,⇑ a

Beijing Nuclear Magnetic Resonance Center, Peking University, Beijing 100871, China College of Chemistry and Molecular Engineering, Peking University, Beijing 100871, China c Department of Physics, University of Guelph, 50 Stone Road East, Guelph, Ontario N1G 2W1, Canada d Biophysics Interdepartmental Group, University of Guelph, 50 Stone Road East, Guelph, Ontario N1G 2W1, Canada b

Edited by Beat Meier and David Neuhaus

a r t i c l e

i n f o

Article history: Received 3 May 2014 Accepted 20 July 2014 Available online 26 July 2014 Keywords: Magic angle spinning solid-state NMR Membrane protein Protein structure Lipid bilayer Multidimensional NMR

a b s t r a c t Membrane proteins mediate many critical functions in cells. Determining their three-dimensional structures in the native lipid environment has been one of the main objectives in structural biology. There are two major NMR methodologies that allow this objective to be accomplished. Oriented sample NMR, which can be applied to membrane proteins that are uniformly aligned in the magnetic field, has been successful in determining the backbone structures of a handful of membrane proteins. Owing to methodological and technological developments, Magic Angle Spinning (MAS) solid-state NMR (ssNMR) spectroscopy has emerged as another major technique for the complete characterization of the structure and dynamics of membrane proteins. First developed on peptides and small microcrystalline proteins, MAS ssNMR has recently been successfully applied to large membrane proteins. In this review we describe recent progress in MAS ssNMR methodologies, which are now available for studies of membrane protein structure determination, and outline a few examples, which highlight the broad capability of ssNMR spectroscopy. Ó 2014 Elsevier B.V. All rights reserved.

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preparation of isotopically labeled membrane protein samples for ssNMR studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. General requirements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Expression of membrane proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.1. Escherichia coli expression systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2. Eukaryotic expression systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.3. Cell-free expression systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Solubilization and reconstitution of membrane proteins for MAS solid-state NMR studies. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Magic angle spinning solid-state NMR techniques for membrane proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Multidimensional spectroscopy for spectroscopic assignments of uniformly 13C,15N labeled proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Interatomic distance measurements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Torsion angle structural restraints . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Paramagnetic relaxation enhancements for structure determination and accelerated data collection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Membrane protein topology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structural and mechanistic insights into membrane proteins by solid-state NMR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. M2 proton channel of influenza A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

⇑ Corresponding author at: Department of Physics, University of Guelph, 50 Stone Road East, Guelph, Ontario N1G 2W1, Canada. E-mail addresses: [email protected] (S. Wang), [email protected] (V. Ladizhansky). http://dx.doi.org/10.1016/j.pnmrs.2014.07.001 0079-6565/Ó 2014 Elsevier B.V. All rights reserved.

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4.2. Microbial rhodopsins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3. Human phospholamban . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4. Potassium channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5. DsbB–DsbA disulfide bond formation complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6. G-protein coupled receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Concluding remarks and perspectives. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Membrane proteins constitute approximately a third of the proteome of living cells [1]. They perform a variety of vital cellular functions including working as receptors that trigger cellular responses to external stimuli, facilitating the transport of ions, metabolites, and other solutes across lipid membranes, catalyzing reactions at the membrane interface, and mediating cell adhesion and growth. Many diseases commonly associated with mutations are linked to the malfunctions of membrane proteins, underscoring their medical importance. More than half of all market drugs are estimated to target membrane proteins. The knowledge of membrane protein structure is one of the key factors required for detailed understanding of their function, and structure determination of membrane proteins has rapidly accelerated in recent years, largely due to the developments of new crystallization methods [2–7]. Especially impressive progress has been achieved in the characterization of G-protein coupled receptors [8–18]. Recent technological and methodological advances in solution NMR have also significantly expanded the applicability of this method to membrane proteins [19–23]. In particular, the introduction of TROSY experiments [24], isotopic labeling strategies [25–31], and advances in novel membrane mimetics, which are compatible with solution NMR [32–35], have enabled structural and dynamic analyses of membrane proteins of both b-barrel and polytopic a-helical folds [36–48]. Despite these achievements, the structural biology studies of membrane proteins significantly lags behind that of their soluble counterparts – as of April 2014 only approximately 1.4% of all protein data bank entries corresponded to the structures of membrane proteins. Furthermore, lipid mimetics, used for both crystallization and solubilization, may distort membrane protein structure [49]. Solid-state NMR (ssNMR) represents a viable and better alternative to X-ray and solution NMR methods in that it can be applied to membrane proteins reconstituted in lipid bilayers of varying lipid composition, over a wide range of pH, temperature and salinity, therefore offering the flexibility to choose optimal sample conditions which closely mimic those of cell membranes. Solid-state NMR can also be applied to the characterization of membrane proteins in native cell membranes [50–55], offering the opportunity to examine the effect of structurally and chemically heterogeneous cellular environments on protein conformation and dynamics. Not less important is that ssNMR is not principally limited by the tumbling rate of a macromolecule, and therefore structural characterization of proteins of large molecular weight is possible. The potential of ssNMR has long been recognized and discussed in the literature [56–61], however many technological and methodological challenges had to be overcome before it could be realized. Historically, the majority of early ssNMR structures have been determined for small a-helical proteins using orientation restraints [62–69]. In this methodology, samples are uniformly aligned, either mechanically on glass plates, or using magnetically aligned bicelles, and 15N–1H dipolar interactions and 15N chemical shift anisotropy are measured to provide restraints on both the helical tilt and local backbone structure. Such measurements generally require multiple selectively labeled samples, and are subjected to

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some practical limitations on the protein size. In contrast, sample preparation for Magic Angle Spinning (MAS) NMR [70,71] does not require alignment, but the experiments are more demanding, as they require relatively fast spinning for efficient averaging of anisotropic dipolar and chemical shift interactions. Recent developments in probe technology and the availability of high magnetic fields have resulted in many additional studies of large polytopic membrane proteins, and many functional and structural insights have been obtained [72–79]. In addition, it was recently demonstrated that orientation restraints can also be collected under MAS conditions using the effect of rotational alignment [80], and this approach resulted in structures for several membrane proteins, including that of G-protein coupled chemokine receptor CXCR1 [81,82]. Our main objective in this article is to highlight recent progress in MAS solid-state NMR studies of membrane proteins which have occurred over the past decade. We emphasize methods and applications to proteins with uniform and/or extensive isotopic labeling, which allows for the complete structural and dynamical characterization of a molecule. For excellent discussions of complementary Oriented Sample (OS) NMR methodology we refer readers to Refs. [83–93]. 2. Preparation of isotopically labeled membrane protein samples for ssNMR studies 2.1. General requirements Successful preparation of protein samples to a large extent defines the success of subsequent ssNMR analysis. Solid-state NMR requires large, milligram quantities of a protein, isotopically enriched with 13C and 15N. For the application of proton detection, membrane proteins may also need to be perdeuterated, with protons reintroduced through back-exchange. The protein has to be natively folded and reconstituted in an environment in which it is stable over the prolonged periods of time required for ssNMR experiments. As the volume of the rotors used for MAS ssNMR is limited, sensitivity considerations often dictate the use of samples with a high protein to lipid ratio. Finally, protein samples must be structurally homogeneous for optimal spectral resolution. Below we review the latest advances in membrane protein sample preparation. For a more detailed discussion of sample preparation strategies we refer readers to recently published articles and reviews [94–96]. 2.2. Expression of membrane proteins 2.2.1. Escherichia coli expression systems E. coli is the most widely used host for heterologous expression and isotopic labeling of proteins for ssNMR studies. Cost-effectiveness, rapid growth rates, high yields, and the availability of a large number of cloning vectors, mutant host strains, and different growth media, are the key advantages of E. coli expression systems. E. coli offers flexibility in choosing simplifying labeling strategies, which are used in ssNMR to improve spectral resolution and to


facilitate chemical shift assignments or structural and dynamic measurements. Uniform 15N and 13C labeling, the most widely used and straightforward labeling strategy, is achieved by expressing a target protein in a minimal medium containing uniformly 13C-labeled glucose as the sole carbon source, and 15N-labeled ammonium sulfate or ammonium chloride as the sole nitrogen source. Multidimensional chemical shift correlation spectroscopy applied to uniformly labeled membrane proteins has resulted in extensive spectroscopic assignments for several of them [79,97–99]. Furthermore, structural restraints can also be obtained on such uniformly labeled samples [100–103]. In applications to large membrane proteins, some degree of spectral simplification is generally required for validation or extension of assignments, and a number of simple labeling approaches have been developed towards this end. Selective amino acid labeling is achieved by adding an excess of isotopically-labeled amino acids to unlabeled growth media, and can be used to create desirable labeling patterns [104–106]. Alternatively, one can reversely label (unlabel) specific residue types through the addition of natural abundance amino acids to a minimal 13C,15Nlabeled culture medium [98,107–110] for spectral simplification. As scrambling can occur between amino acids with shared biosynthetic pathways, the efficiency of selective labeling/unlabeling can be enhanced through the use of auxotrophic strains [111,112]. Isotopic dilution, which is commonly used to simplify the labeling patterns of amino acids and offers a number of practical advantages, is achieved by growing a protein on [1,3-13C] glycerol, [2-13C] glycerol, [1-13C], or [2-13C] glucose as the sole carbon source [107,113–116]. This labeling scheme reduces the crowdedness of NMR spectra and improves carbon resolution by removing most one bond 13C–13C J-couplings. The spectral simplification and resolution enhancement afforded by such labeling is especially noticeable in the aromatic region, and has been shown to facilitate spectroscopic assignments of aromatic side chains [117,118]. Additionally, dilution of 13C spins in a protein reduces dipolar truncation effects [119], simplifies the polarization transfer pathways, and facilitates measurements of long-range distance restraints, which are critical for structural determination [114,120,121]. E. coli is a versatile expression system and will remain an attractive choice for the production of isotopically labeled samples of membrane proteins, especially prokaryotic ones. Over-expression of eukaryotic membrane proteins is much more challenging in E. coli, especially for systems which require posttranslational modifications or complicated eukaryotic machinery for proper folding and insertion into the membrane. Often, eukaryotic membrane proteins express into inclusion bodies in E. coli, and require refolding protocols. To this end, Huster and co-workers have published an in vitro refolding protocol for the G-protein coupled neuropeptide Y2 receptor, and have demonstrated an improvement in 15N dispersion of the receptor upon reconstitution in lipids [122,123]. Gawrisch and co-workers have also developed a refolding protocol for the human Cannabinoid Type 2 receptor [124]. Despite these promising results, the spectral resolution obtained for these two GPCRs is not yet on par with that available for other membrane proteins, and further optimization is required before detailed, site-specific analysis could be carried out in these systems. On the other hand, the successful structure determination of CXCR1 [82], which was expressed in inclusion bodies in E. coli and subsequently refolded [125], demonstrated the potential utility of E. coli for the production of eukaryotic proteins. 2.2.2. Eukaryotic expression systems Yeast is the simplest eukaryotic host for heterologous protein expression, and represents a viable alternative for the production of eukaryotic proteins. Pichia pastoris and Saccharomyces cerevisiae are the two commonly used yeast hosts for recombinant protein

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expression for X-ray crystallography and NMR studies [126,127]. In particular, the potential of methylotrophic yeast P. pastoris to express isotopically labeled proteins has been extensively investigated, and protocols for the production of isotopically labeled samples using 13C-labeled methanol as a primary carbon source are available [128–132]. Brown and co-workers have recently demonstrated the utility of this host for the production of isotopically 15 13 N, C-labeled membrane proteins for ssNMR. Leptosphaeria rhodopsin from Leptosphaeria maculans was expressed and isotopically labeled at high levels and resulted in well resolved solid state NMR spectra suitable for structural analysis [133]. A similar protocol was optimized for the expression of the water channel human Aquaporin-1, and resulted in highly resolved ssNMR spectra [134]. Furthermore, many GPCRs could be expressed in these yeast cells [135], and would, in principle, be amenable to isotopic labeling for ssNMR. The utility of P. pastoris is further enhanced by the availability of auxotrophic strains [136], and deuteration protocols [130]. Higher eukaryotic expression systems, e.g., transfected mammalian cells and baculovirus-infected insect cells, contain the necessary cellular machinery and offer flexibility for native folding and targeting, and for post-translational modifications. Although there has been some degree of success in the development of methods for isotopic labeling [137–139], high cost, incompleteness of labeling, and low yields limit the applicability of higher eukaryotic expression systems for complete structure determination by NMR. Residue specific labeling is more successful, and has been used for the characterization of bovine rhodopsin [140–146]. 2.2.3. Cell-free expression systems In vitro cell free expression systems use the coupled transcription and translation reaction to produce isotopically labeled proteins. They lack the amino acid biosynthesis systems, and therefore require the addition of labeled amino acids to the reaction mixture. This offers a number of advantages in the choice of labeling strategies, as amino acids can be labeled selectively without scrambling (as there are no enzymes responsible for the amino acid synthesis), and there are protocols available for combinatorial labeling [147–152], and for stereo-array isotopic labeling (SAIL) [153,154]. On the other hand, the addition of labeled amino acids may add to the overall cost of the protein sample. While the most extensively used cell free systems use bacterial extracts, systems have also been developed which use alternative cell lines such as wheat germ and insect cells [155,156]. An important consideration, which defines the effectiveness of cell free expression systems for membrane protein production, is whether they can produce correctly folded proteins. In this respect, the presence of mild detergents, or even lipids, in the reaction reduces aggregation and insolubility, and increases the expression yields of correctly folded proteins [157–160]. 2.3. Solubilization and reconstitution of membrane proteins for MAS solid-state NMR studies Once a membrane protein is expressed at an acceptable level, it has to be isolated and reconstituted in a membrane mimetic environment: proteins must be extracted from the cell membrane using detergents, and reconstituted back into a model lipid membrane following the purification step. The choice of solubilizing detergent depends on whether the protein is expressed into inclusion bodies, or properly inserted into the host cell membrane. For the latter, mild detergents preserving the native protein fold can be used for solubilization. Typical choices of detergents are DDM, OG, DPC, or Triton X-100 (Fig. 1A). In each case the choice of detergent is protein specific and requires optimization to obtain complete solubilization and to avoid protein aggregation and misfolding. For membrane proteins expressed into inclusion bodies,

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Fig. 1. (A) Common detergents used for membrane protein solubilization from cell membranes. (B) Typical lipids used for reconstitution of membrane proteins for solid state NMR. They have different headgroups, acyl chains, and number of unsaturated bonds.

solubilization requires harsh detergents such as EmpigenÒ BB or SDS during the initial stage, often in combination with the use of denaturants [36,37,161]. Subsequent refolding of membrane

proteins solubilized from inclusion bodies is a challenging task, especially for large membrane proteins, and requires the careful optimization of experimental conditions [95,122,125].


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Fig. 2. Magic angle spinning carbon–carbon correlation spectra of various classes of membrane proteins. (A) Spectrum of YadA collected at 900 MHz. Protein sample was prepared in a microcrystalline form. Reprinted by permission from Macmillan Publishers Ltd.: Scientific Reports, 2, S.A. Shahid, S. Markovic, D. Linke, B.J. van Rossum, ‘‘Assignment and secondary structure of the YadA membrane protein by solid-state MAS NMR’’, (2012) 803. Copyright 2012. (B) 13C–13C correlation spectrum of lipidreconstituted chimeric potassium channel KcsA-Kv1.3 collected at 800 MHz. Adapted with permission from J. Am. Chem. Soc., 130, R. Schneider, C. Ader, A. Lange, K. Giller, S. Hornig, O. Pongs, S. Becker, M. Baldus, ‘‘Solid-state NMR spectroscopy applied to a chimeric potassium channel in lipid bilayers’’ (2008) 7427–7435. Copyright 2008 American Chemical Society. (C) 13C–13C correlation spectrum of DsbB collected at 750 MHz. Protein is prepared in a precipitated state with endogenous lipids present. Reprinted from Journal of Molecular Biology, 425, M. Tang, A.E. Nesbitt, L.J. Sperling, D.A. Berthold, C.D. Schwieters, R.B. Gennis, C.M. Rienstra, ‘‘Structure of the disulfide bond generating membrane protein DsbB in the lipid bilayer’’ (2013) 1670–1682. Copyright 2013, with permission from Elsevier. (D) 13C–13C correlation spectrum of lipid-reconstituted Anabaena Sensory Rhodopsin collected at 800 MHz. Reproduced with permission from Angewandte Chemie Int Ed., ‘‘Conformation of a seven-helical transmembrane photosensor in the lipid environment’’ 50 (2011) 1302–1305. Copyright 2011 John Wiley & Sons, Inc. (E) 13C–13C correlation spectrum of water channel human Aquaporin 1 collected at 800 MHz. Protein was reconstituted in lipids. Reprinted from Journal of Biomolecular NMR, S. Emami, Y. Fan, R. Munro, V. Ladizhansky, L.S. Brown, ‘‘Yeastexpressed human membrane protein aquaporin-1 yields excellent resolution of solid-state MAS NMR spectra’’ 55 (2013) 147–155. Reproduced with kind permission from Springer Science and Business Media.

Membrane protein structure and function is often sensitive to the environment [49], and solid-state NMR offers great flexibility in choosing the appropriate sample conditions. The native-like lipid bilayer represents the most attractive environment for membrane protein sample preparation. For lipid reconstitution, protein solubilized in detergent is mixed with lipids, which results in the formation of a mixed detergent/lipid/protein micelle. Detergent can be removed by dialysis, or using polystyrene resins such as Bio-beadsÒ SM [162]. There are a variety of lipids that can be used for membrane protein reconstitution (Fig. 1B). In each case, the lipid composition and the protein-to-lipid ratio should represent a compromise between physiological relevance and experimental tractability. DMPC/DMPA binary mixtures were successfully used for microbial rhodopsins [99,110], as the presence of a small proportion of anionic DMPA (10%) is important for the rhodopsins’ function and orientation [163]. A binary mixture of Egg PC/Brain PS resulted in spectra of high quality for human Aquaporin 1. A mixture of zwitterionic DOPE and anionic DOPS was used for reconstitution of the KcsA potassium channel [164]. There are a number of recently published protocols detailing the sample preparation procedures for ssNMR studies and we refer readers to these excellent papers for more details [95,96,133,165,166]. While reconstituting membrane proteins in lipids is preferred from the physiological relevance standpoint, in some cases the

microcrystalline environment can also be used for membrane protein ssNMR studies [73,77,97]. Higher protein density can be achieved in microcrystals, and a breadth of biochemical conditions can be applied for studies of protein function. On the other hand, protein structures obtained from such samples are liable to crystallization artifacts similar to those seen in large crystals. Fig. 2 shows some representative 13C–13C correlation spectra of membrane proteins of different classes [117,118], prepared in different environments.

3. Magic angle spinning solid-state NMR techniques for membrane proteins 3.1. Multidimensional spectroscopy for spectroscopic assignments of uniformly 13C,15N labeled proteins Obtaining resonance assignments for a protein is a prerequisite for the site-specific analysis of its structure, dynamics and function. The choice of assignment methodologies and their success depend on a number of factors such as molecular weight, structural homogeneity, and overall mobility of the protein system of interest. To date, the majority of membrane protein studies use carbon detection, and 13C-detected assignment strategies have been

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Fig. 3. An example of solid-state NMR assignment strategies. (A) The NCOCX experiment (top panel) correlates chemical shifts of nitrogen N[i], and CO[i-1] and CX[i-1] of the preceding residue. The CANCO experiment (middle panel) establishes interresidue correlations between the backbone CA[i], N[i] and CO[i-1] atoms. The NCACX experiment (bottom panel) correlates N[i], with CA[i] and CX[i]. Extended spin systems shown in (B) can be built by matching the backbone chemical shifts detected in these experiments. (B) Two extended spin systems can be combined into a contiguous fragment by matching shifts of backbone and side chain atoms shown in dashed boxes. Reprinted from Methods of Molecular Biology, L. Shi and V. Ladizhansky ‘‘Magic angle spinning solid-state NMR experiments for structural characterization of proteins’’, 895 (2012) 153–165. Reproduced with kind permission from Springer Science and Business Media.

established. The general approach in solids is similar to that used in solution NMR and relies on recording multiple multidimensional correlation spectra which can be related to one another through a series of shared correlations, and used to construct the backbone walk. While for smaller proteins with fewer resonances twodimensional spectroscopy appears to be sufficient to assign most resonances (e.g., microcrystalline a-spectrin SH3 domain [167], GB1 [168]), obtaining reliable assignments in proteins of higher molecular weight requires three-dimensional correlation spectroscopy, and often needs to be combined with simplifying labeling strategies. Fig. 3 shows one of the common assignment strategies, which relies on three basic 3D correlation experiments: (i) CANCO experiment establishes correlation between the three backbone atoms, CA[i], N[i], and CO[i-1]; (ii) NCACX experiment provides intraresidue correlation between the backbone atoms N[i], CA[i] and CX[i] (CX denotes both the backbone and side chain atoms, CO, CB, CG, etc.); (iii) NCOCX experiment correlates the backbone atom N[i] to the carbonyl CO[i-1] and CX[i-1] of the preceding residue. These three experiments can be used to construct the backbone walk as illustrated in Fig. 3, and at the same time identify residue type through correlations of the side chain resonances. Additional correlations for the side chain atoms can often be derived from two dimensional carbon–carbon correlation spectra. One of the potential complications of the NCOCX, NCACX and CANCO strategy is that two of the experiments rely on the carbonyl chemical shift dispersion, which is typically poor and results in high degree of spectral overlap in the NCO plane. Meier, Böckmann and co-workers have proposed a complementary strategy relying on NCACB, N(CO)CACB and CAN(CO)CA experiments, which replace the poorly dispersed CO frequencies with those of CB atoms [169]. Alternatively, the resolution of the NCO plane can be substantially improved by employing 2-13C and 1,3-13C glycerol labeling schemes, which reduce the number of resonances and enhance the CO resolution by suppressing the one bond CA-CO J-couplings. An additional benefit from glycerol-labeled samples is that they facilitate the assignment of the aromatic side chains, many of which are poorly resolved in uniformly labeled samples because

of the high degree of spectral overlap and broad lines [117,118]. Although much less sensitive, four-dimensional spectroscopy has been demonstrated to result in a substantial increase in spectral resolution. Originally demonstrated in the small microcrystalline GB1 protein [170], 4D chemical shift correlation experiments have been used in membrane proteins as well [79,97,108]. The solvent exposed loops, turns, and termini of membrane proteins, are frequently subjected to restricted motions of varying time scales and amplitudes. Overall, these motions partially average internuclear dipolar couplings and reduce the efficiency of the dipolar-driven polarization transfer steps in correlation experiments. The detection and unambiguous assignment of such fragments of intermediate mobility remain a challenge. Although alternative through-bond multidimensional correlation methods have been demonstrated on microcrystalline proteins [171–173], their applications to membrane proteins remain limited to some extreme cases when large amplitude motions are present. Baldus and co-workers have used INEPT based experiments to assign resonances of the C-terminal mobile tail of phospholamban [174]. Similar methodology was applied to assign mobile fragments of a 7TM protein sensory rhodopsin II from Natronomonas pharaonis [109]. More economical versions of the scalar-based heteronuclear chemical shift correlation experiments were developed by Zhong et al. [175]. The constant–time implementation of the 15N/13C INEPT polarization transfer steps and the chemical shift evolution periods resulted in substantial sensitivity improvements, and permitted assignments of the mobile fragments of peripheral membrane-associated myelin basic protein to be made. 3.2. Interatomic distance measurements Interatomic distances are the main source of structural information in both solution and solid state NMR. In the solid state, the internuclear dipolar interactions averaged out by magic angle spinning can be reintroduced through the application of radiofrequency pulses, an approach dubbed dipolar recoupling [176,119]. Dipolar recoupling allows accurate distances to be


measured in selectively labeled proteins [177], but meets one principal complication in uniformly 13C labeled proteins: the dipolar Hamiltonians corresponding to structurally constraining weak dipolar couplings are truncated by the strong ones, rendering long distance measurements inefficient [119]. In most cases, a combination of spin dilution through sparse labeling schemes, and/or second-order dipolar recoupling methods, which are less sensitive to dipolar truncation effects, is required to facilitate internuclear distance measurements in proteins. At slow to moderate spinning rates proton-driven spin diffusion (PDSD) [178] or dipolar assisted rotational resonance (DARR) [179,180] have been used extensively to probe internuclear distance restraints [76–78,114,120], while at faster rates where DARR or PDSD become inefficient, a new generation of second order recoupling techniques have been developed for both homonuclear [181–184] and heteronuclear [185–188] distance measurements. Interproton distances are usually measured in combination with carbon detection, [77,78,100,101,189,190], and represent another rich source of structural information. Frequency selective recoupling methods are a useful alternative that can be used for accurate distance measurements in uniformly 13 C labeled proteins. Among them, rotational resonance (R2) [191] and related techniques have been demonstrated to result in structurally constraining distances in peptides and proteins [177,192]. In particular, Griffin and co-workers have proposed a rotational resonance width experiment [193], in which the internuclear carbon–carbon dipolar interactions are recoupled in a constant–time manner, and the polarization transfer is recorded as a function of spinning frequency, which is varied across the rotational resonance condition. The resulting resonance shapes can be fitted as a function of the dipolar coupling and the zero-quantum relaxation rate, and allow for the unambiguous extraction of internuclear distances [121,194]. Furthermore, using reduced decoupling during the R2 recoupling causes homogeneous broadening of the R2 matching condition, and results in band-selective (rather than frequencyselective) polarization transfer between the carbonyls and aliphatic side chains simultaneously [103]. This technique, named homogeneously broadened rotational resonance (HBR2), has been used to study Anabaena Sensory Rhodopsin, and provided more than two dozens of medium and long range restraints (|i j| > 3) and almost two hundred short range distance restraints ((|i j| 6 3) for structure determination [78]. There are a variety of techniques available for heteronuclear distance measurements. The most widely used ones in the context of a complete structure determination are based on rotational echo double resonance [195] or transferred echo double resonance [196]. These methods were combined with multidimensional spectroscopy applications in uniformly 13C, 15N-labeled peptides and proteins [197,198], and have also been used to probe protein interfaces [199].

3.3. Torsion angle structural restraints The addition of torsion angles to the list of structural restraints used in the structure calculation protocol greatly improves the quality of NMR structures. Torsion angles can be estimated from chemical shifts of backbone atoms using available software tools such as TALOS+ [200]. Alternatively, torsion angles can be probed directly using recoupling methods by correlating suitable anisotropic interactions. A large library of such correlation experiments have been developed for probing both / and w backbone torsion angles. For example, the /i (Ci-10 –Ni–Cia–Ci0 ) backbone torsion angle can be constrained by correlating 15Ni–1Hi and 13Ci–1Hi dipolar tensors [201,202], or by correlating the relative orientation of the 15N chemical shift anisotropy and 13Ci–1Hi dipolar tensors [203].

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The wi backbone torsion angle can be obtained by correlating relative orientations of the 15Ni+1–13C0 i and 15Ni–13Cia dipolar tensors [204,205], 13Ci–1Hi and 15Ni+1–1Hi+1 [206] tensors, which are sensitive to the variation of w in the range corresponding to beta secondary structure, or 15Ni+1–13C0 i and 13Cia–1Hi dipolar tensors [207], which are more suitable for applications in alphahelical proteins. There is also a series of methods, which explores the dependence of the relative orientation of the CSA tensors and dipolar interactions [208–210]. These tensor correlation methods can be combined with multidimensional chemical shift correlation spectroscopy for site-specific resolution and have led to multiple restraints on protein structure [206,211–214]. In particular, the latter studies on GB1 have demonstrated that the dipolar correlation methods and chemical shift tensor analysis can substantially improve structural resolution, and are a powerful tool for structure refinement [213,214]. Dipolar correlation methods have been used in membrane proteins to answer mechanistic questions related to protein function. Chromophore distortions during the photocycle of bacteriorhodospin were probed using an HCCH [215] dipolar correlation experiment. These measurements indicated that the all-trans retinal deviates from planarity around the C14–C15 bond in light adapted bR, and this deviation increases in the M intermediate state [216]. Similar measurements were performed on bovine rhodopsin, a light sensitive receptor responsible for vision under low-light conditions. The H–C10–C11–H torsional angle in the photo-intermediate Meta I state, which is a precursor for the Meta II active state, was found to be consistent with the all-trans conformation of retinal [217]. 3.4. Paramagnetic relaxation enhancements for structure determination and accelerated data collection Although the use of paramagnetic ssNMR is limited in membrane proteins at present, employing paramagnetic tags, either for collecting structural restraints, accelerated data collection, or probing protein topology, would be notably beneficial. Below we briefly review the current state of paramagnetic ssNMR. Obtaining long-range distance information is critical for determining correct protein structures. Long-range restraints are sparse in ssNMR, as typical detectable carbon–carbon and carbon–nitrogen distances do not exceed 5–6 Å. The electron–nuclear spin couplings are much stronger and can be used to collect long-range structural distance restraints. The use of paramagnetic tags, either endogenously present in a protein or introduced exogenously, has become an established tool in solution NMR [218–220], and is also making its way into the biomolecular ssNMR toolbox [221–223]. Paramagnetic centers can be introduced into a protein through metal binding sites, if such sites are endogenously present, or through chemical modification of reactive cysteine side chains. Cysteines, either endogenously present or introduced through mutagenesis, can be reacted with cysteine-reactive paramagnetic tags, most commonly the nitroxide radical MTSL, or EDTA loaded with paramagnetic metals. The effect induced by the paramagnetic center depends on its properties: (i) paramagnet can result in a Fermi contact shift if there is an electron spin density present at the position of a nucleus; (ii) Pseudocontact shifts (PCSs) are induced by paramagnetic centers with significantly anisotropic gtensors (e.g., Co2+, Yb3+, etc.); (iii) Paramagnetic tags with both anisotropic and isotropic g-tensors (e.g., nitroxide, Cu2+, Mn2+, etc.) result in enhanced NMR relaxation rates (paramagnetic relaxation enhancement, PRE) of nearby nuclear spins. Paramagnetic centers with short electron relaxation times and high anisotropic electronic magnetic susceptibility lead to significant PCSs, which depend on the electron–nucleus distance and on the orientation of the electron–nucleus vector in the principal

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axis frame of the magnetic susceptibility tensor. Many paramagnetic transition metal ions and lanthanide ions, except Cu2+, Mn2+ and Gd3+, could induce large PCSs. Although PCS effects have not yet been used in ssNMR studies of membrane proteins, their potential utility for studies of protein structure, protein–protein, and protein–ligand interactions has been demonstrated. PCSs induced by a Fe3+ ion bound to the Cytochrome P450 were utilized to characterize the conformational changes occurring in the enzyme upon ligand binding [224]. The structure of the Co2+ substituted matrix metalloproteinase 12 (CoMMP-12) [225] has been determined using PCSs from endogenous paramagnetic ions. Structural PCSs can also be obtained from exogenous paramagnetic ions introduced via paramagnetic ion-chelated tags, which can be covalently attached to cysteines introduced by sitedirected mutagenesis [226]. PSCs perturb the cross peak positions in the spectra, and additional chemical shift assignments may be necessary for data interpretation. In contrast, paramagnetic centers with isotropic g-tensors result in relaxation enhancement only, without affecting the peak frequencies. The PRE effect in solids is inversely proportional to the sixth power of the electron–nucleus distance [222], and can be used to obtain easily interpretable structural restraints. Both longitudinal R1 and transverse R2 relaxation can be enhanced, depending on the electron relaxation properties of a paramagnetic center. The nitroxide radical MTSL has a typical T1e relaxation time of about 100 ns, and results in transverse R2 relaxation enhancement, but to a much lesser extent affects longitudinal relaxation. In practice, R2 PREs induced by the MTSL label result in significant signal attenuation for nuclear spins within approximately a 15 Å radius, providing a quick way to collect semi-quantitative structural restraints [227]. More accurate quantification of transverse PREs in solids is complicated by a number of factors, in particular by rapid interproton spin diffusion. Jaroniec and co-workers have recently demonstrated an alternative way to measure accurate electron–nuclear distances. Cu2+ has a short T1e in the nanosecond range at high magnetic fields and at near ambient temperatures, and yields large longitudinal PREs, but produces negligible line broadening effects. Paramagnetic enhancement of 15N R1 rates can be accurately quantified and have led to electron–nuclear distance restrains in the range from 13 to 24 Å in the GB1 protein [228]. It was later demonstrated that PREs collected from Cu2+ are sufficient for the determination of a global fold of GB1 [229]. 15N PRE effects induced by Cu2+ were also used for structure determination of the Cu–Zn superoxide dismutase structure [230], and for the detection of copper binding sites in the M2 proton channel [231]. Owing to their long-range nature, both PCSs and PREs can be used for the characterization of intermolecular interactions, crystal packing, and oligomerization, while inter- and intramolecular paramagnetic effects can be disentangled through the dilution of paramagnetically labeled protein in a diamagnetic background [225,232]. In this way, PCSs allowed for the characterization of intermolecular packing and determination of the crystal structure of CoMMP-12 [233]. MTSL nitroxide spin labels have also been successfully used for the characterization of the oligomerization interface of a trimeric 7TM membrane protein Anabaena Sensory Rhodopsin (ASR) [234]. In this study, intra-monomer PREs partially defining relative positions of some of the helices, and intermonomer PREs defining the oligomer interface, were obtained from a single MTSL label attached to a genetically engineered cysteine [234]. The addition of paramagnetic ions into a protein sample results in the shortening of proton relaxation times, and can be used for the rapid acquisition of solid-state NMR data. In combination with fast magic angle spinning and low power decoupling [235], very fast recycle delays can be afforded without causing significant

sample or probe heating [236,237]. Paramagnetic tags that preferentially enhance longitudinal relaxation can be either covalently attached to the protein [238,239], or exogenously added to buffer [239,240], or chelated to lipids [240–242]. The most frequently used tag, EDTA:Cu2+, allows for an approximate ten-fold R1 relaxation rate increase, while the recently tested Gd3+–DOTA complexes can shorten relaxation times by almost two orders of magnitude, as was recently demonstrated in a 7TM protein proteorhodopsin [243]. 3.5. Membrane protein topology Transmembrane proteins contain both large hydrophobic domains which are embedded into a lipid bilayer and are often inaccessible to solvent, and solvent exposed regions, e.g., loops and turns, flanks of helices, or polar cavities inside the protein. Paramagnetic relaxation enhancement effects induced by paramagnetic ions that preferentially partition into solution or lipids can be used to identify residues that are close to the interface or located in the hydrophobic core, respectively. The solvent exposed regions can also be probed by hydrogen–deuterium exchange, or using water-edited experiments, in which the nuclei at the protein–water interface are excited from the protons of water. In the latter approach, however, the interproton spin diffusion results in rapid redistribution of signal across the protein, and this may limit the selectivity of the protein–solvent interface detection. Mn2+ ions added in the buffer partition into the polar solvent, selectively enhance spin–spin lattice relaxation and suppress the spectral lines of solvent exposed residues. This approach was used to study the depth of membrane insertion of protein fragments of bacteriorhodopsin [244] and of the antimicrobial peptide protegrin-1 [245]. Alternatively, dioxygen preferentially partitions towards the hydrophobic portion of the lipid membrane, perturbs chemical shifts, enhances spin–lattice relaxation, and can be used to characterize the depth of immersion of membrane peptides and proteins [246]. Modification of lipids with paramagnetic groups can also be used to characterize the depth of penetration of membrane proteins into lipid bilayers [247]. Hydrogen–deuterium exchange provides a simple way to probe the solvent-accessible surface of membrane proteins. In the simplest implementation, the amide protons of the solvent exposed and exchangeable residues of a membrane protein incubated in D2O, are replaced with deuterons, and this results in the disappearance of the signals of these residues in 2D NCA or 3D NCACX/ NCOCX experiments, thus providing a site-specific view of the protein–water interface, and also reporting on the strengths of intrahelical hydrogen bonds [248,99]. This method has also been successfully used to indirectly detect functional intermediate states, for example, those occurring in the photocycle of ASR [249]. More recently, H/D exchange was used to probe the solvent accessibility of the DAGK enzyme reconstituted in E. coli lipids [250]. Water-edited experiments provide an alternative method for the identification of protein fragments exposed to solvent. Coherence lifetimes of the protons within a protein are much shorter than those of mobile species, e.g., water and lipids, and thus protein bound protons can be selectively suppressed using a transverse relaxation filter. Protein protons located at or near the interface can be selectively excited from the long lived water or lipid signals, either through the chemical exchange mechanism [251,252], or via through-space transfer. In one of the first applications of this idea, Harbison et al. monitored the kinetics of magnetization transfer from solvent water to a single site in bacteriorhodospin (bR), and obtained evidence that the Schiff base proton of bR undergoes direct chemical exchange with the bulk water [251]. This approach was later used to study protein–water


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Fig. 4. Structures of intrinsic membrane proteins determined either by ssNMR, or by combining solution and ssNMR, or X-ray and ssNMR restraints. Protein name, molecular weight, used membrane mimetic, method by which structure was determined, and PDB ID code are given below each structure.

interactions in crystals [253–256], and for the identification of the water exposed residues of sensory rhodopsin SRII [109]. Water-edited experiments have also been used to monitor changes in the water–protein interface of the chimeric potassium channel KcsA-Kv1.3, and to identify structural rearrangements occurring in response to changes of pH [257]. 4. Structural and mechanistic insights into membrane proteins by solid-state NMR

ssNMR restraints and resulted in improved structural resolution [76,259]. Additionally, many studies have focused on membrane protein function, through detailed characterization of the conformational and dynamic changes under various stimuli, e.g., changes in pH, light activation and interactions with ligands or other proteins. Below we highlight a few examples of membrane protein studies of the past decade. 4.1. M2 proton channel of influenza A

While the majority of early ssNMR studies focussed on small membrane proteins or on proteins with selective isotopic labels, technological and methodological advancements over the past decade have enabled many applications of ssNMR to the detailed structural analysis of large polytopic proteins. As of April 2014, there were 23 entries for structures of membrane proteins determined by ssNMR techniques (Fig. 4). The structures of six large membrane proteins have been determined by ssNMR within the last two years, including the pentameric human phospholamban [258], the chemokine receptor CXCR1 [82], Yersinia adhesin A (YadA) [77] (a b-sheet rich outer membrane protein), and Anabaena Sensory Rhodopsin [78]. Additionally, structures of DsbB, and of DsbB–DsbA complex were solved by combining X-ray data and

The M2 proton channel protein is found in the viral envelope of the influenza A virus. Its main function is to conduct protons across the viral membrane, performing the critical step in acidification of the interior of the virus, and to modulate the pH of the trans-Golgi network later in the life cycle [260,261]. M2 is a 97-residue long peptide, which consists of an N-terminal domain (residues 1–23), transmembrane (TM) domain (residues 24–46), and C-terminal domain (residues 47–97). It exists as a tetramer in the native state, and forms a pH activated proton channel, which can be inhibited by the antiviral drugs amantadine and rimantadine [262], as well as by Cu2+ ions [263]. As a proven pharmacological target, M2 has been extensively studied by a variety of structural and

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Fig. 5. Solid-state NMR structure of M2 and its mechanism of proton conduction. (A) OS ssNMR structure of the M2CD tetramer. (B) Constrictions at Val27 and Trp41, and His37 tetrad in closed state. From Science 330, ‘‘Insight into the mechanism of the influenza A proton channel from a structure in a lipid bilayer’’, M. Sharma, M. Yi, H. Dong, H. Qin, E. Peterson, D.D. Busath, H.X. Zhou, T.A. Cross, 509–512 (2010). Reprinted with permission from AAAS, and modified with permission from the authors. (C) Structure of the His37 tetrad, responsible for pH sensing and proton selectivity, and proposed water orientations. From Science 330, ‘‘Mechanisms of proton conduction and gating in influenza M2 proton channels from solid-state NMR’’, F. Hu, W. Luo, M. Hong, 505–508 (2010). Reprinted with permission from AAAS, and modified with permission from the authors.

Fig. 6. Amt–M2 interactions. (A) 13C–2H REDOR distance measurements at two different Amt:channel ratios. Both reference spectra (S0), REDOR dephased spectra (S) and difference spectra (DS) are shown. At a low Amt:channel ratio only internal residues show REDOR dephasing, which is consistent with a primary binding site being located inside the helical bundle. At a high Amt:channel ratio secondary binding sites result in the dephasing of additional residues. (B) Sideview and (C) topview of the solid-state NMR structure of Amt-bound M2. Reprinted by permission from Macmillan Publishers Ltd.: Nature, ‘‘Structure of the amantadine binding site of influenza M2 proton channels in lipid bilayers’’, S.D. Cady, K. Schmidt-Rohr, J. Wang, C.S. Soto, W.F. Degrado, M. Hong, 463, 689–692, Copyright 2010.

biophysical methods, including investigations by ssNMR, aiming to elucidate the mechanism of proton conduction by M2, and the mechanisms of its inhibition by drugs. Early ssNMR backbone structures were obtained for the truncated version of the protein, which represents the TM domain (M2TM), using orientation restraints from PISEMA experiments on M2 samples reconstituted in DMPC lipids, and aligned on glass plates [66,264–266]. The PISEMA restraints helped to determine

the TM helix orientation with respect to the bilayer [66]. Additional REDOR measurements between the selectively labeled 15 Np of His37 and 13Cc of Trp41 provided an intermonomer distance restraint on the tetrameric structure [266]. Several structures for longer constructs consisting of the TM helix and the C-terminus (M2CD), which is important for tetramer stabilization and native-like proton conductance [267], have recently been determined using ssNMR. In the solid state NMR


structure determined at pH 7.5, which promotes the closed channel conformation, the monomer consists of a transmembrane helix with a glycine kink, and an amphipathic helix in the C-terminus, which is positioned nearly parallel to the bilayer [268]. The four monomers are arranged in a tetramer stabilized by contacts between nonpolar residues located in the amphipathic helices (Fig. 5). The key residues involved in H+ transport are located in the transmembrane domain: Val27 and Trp41 are responsible for gating at the entrance on the extracellular side and the exit on the cytoplasmic side, respectively, and His37 represents a pH sensor, and is responsible for proton selectivity. The four His37 residues face the core of the helical bundle and compose a histidine tetrad stabilized by strong interhelical Ne2–H–Nd1 hydrogen bonds formed between two adjacent histidine residues, which results in a imidazole–imidazolium dimer structure [268,269] (Fig. 5). A dimer of dimers arrangement was also observed in the full-length M2 proton channel both in synthetic lipid bilayers and in native E. coli membranes [270], and in the drug-resistant S31N mutant of the M2 conducting domain [271]. Investigations of the pH dependence of His37 protonation states, rotameric conformation, and sidechain dynamics in the TM fragment of M2 have shed light on the mechanism of proton conduction. In the closed conformation of the channel at high pH of 8.5, isotropic chemical shifts of His37 revealed neutral imidazoles, with the Ne2 protonated s tautomer and the Nd1 protonated p tautomer existing in a ratio of 3:1 [272–274]. In the open conformation at low pH, the histidines form double protonated cationic states. Measurements of dipolar couplings were consistent with the two site jump of imidazolium rings, suggesting that the His ring reorientation is directly involved in proton transport, overall supporting the shuttle model of proton conductance (Fig. 5C) [273]. As M2 is a proven pharmacological target, the mechanism of interaction between M2 and various drugs and inhibitors has attracted considerable attention. The antiviral drug Amantadine

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(Amt) binds the channel, causing in the formation of a more compact structure of the TM domain, and blocking proton transport [74,275]. Carbon–deuterium REDOR distance measurements between perdeuterated Amt and the protein suggested the presence of two binding sites with different affinities [74]. The primary binding site, occupied at low drug/protein ratio, is located in the cavity between Val27 and His37 in the core of the helix bundle (Fig. 6) and its location remains unaffected in the longer M2CD construct [276]. Amt bound in this cavity undergoes rapid rotation about an axis tilted at 13° with respect to the axis of the channel as revealed by line shape analysis of 2H NMR spectra. When Amt is in excess, it can also associate nonspecifically on the surface of the protein [74]. Another common antiflu drug, rimantadine, binds M2CD specifically at a similar site to the amantadine site, and induces a large-scale structural rearrangement of the entire TM region, supporting the allosteric mechanism of inhibition [277]. 4.2. Microbial rhodopsins Microbial rhodopsins are a diverse group of photosensitive membrane proteins, found in archaea, eubacteria, and unicellular eukaryote organisms [278]. They can perform diverse ion-pumping or sensorial functions in response to light excitation. A typical microbial rhodopsin consists of a seven-helical apoprotein and an all-trans retinal, a light-sensitive pigment, covalently bound to a lysyl residue in the middle of seventh helix (helix G) (Fig. 7). The light-driven proton pump, bacteriorhodopsin (bR) from Halobacterium salinarum is the best-known example of a microbial rhodopsin. It has been studied extensively since its discovery in the 1970s, serving as a prototypical transmembrane ion pump, an early model for G protein-coupled receptors, as well as a testing ground for many biophysical techniques. A wealth of information is available on the bR structure and on the details of its photocycle, making it one of the best characterized membrane proteins. Solid-state NMR has also been extensively used to characterize the biochemical and dynamic properties of bR, as well as structural

Fig. 7. Microbial rhodopsins studied by ssNMR. All proteins are shown as seven-helical bundles. The retinal chromophore shown in red is covalently linked to a conserved lysine residue in the seventh helix. Colors shown are approximately the color of the pigments. Bacteriorhodopsin and proteorhodopsin are proton pumps. Sensory Rhodopsin II interacts with its membrane-embedded transducer (HtrII), and initiates a photophobic response to blue/green light. Anabaena Sensory Rhodopsin interacts with a soluble transducer (ASRT), and is believed to be responsible for the regulation of genes of several light harvesting proteins.

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Fig. 8. Proton-detected spectroscopy in PR. (A) An example of a 2D plane of a three-dimensional CONH correlation spectrum with assigned residues shown. Spectrum was collected at 800 MHz. (B) 3D model of PR based on the homology modeling using BR structure (PDB ID: 1C3W) as a template. Colored residues (blue and green) have been assigned by solid state NMR. Blue residues are observed in proton-detected experiments and have exchangeable amides. In particular, the inner side of helix G is exchangeable. Reprinted from the Journal of the American Chemical Society, 133 ‘‘Proton-detected solid-state NMR reveals intramembrane polar networks in a seven-helical transmembrane protein proteorhodopsin’’, M.E. Ward, L. Shi, E. Lake, S. Krishnamurthy, H. Hutchins, L.S. Brown, V. Ladizhansky, 17434–17443 (2011). Copyright 2011, American Chemical Society.

changes occurring during its photocycle [177,216,279–288]. More recently, the Dynamic Nuclear Polarization (DNP) [289] technique was used for the detection of weakly populated reaction intermediates of bR, which are invisible to other methods. DNP-enhanced ssNMR has provided the first NMR observation of 15N Schiff base shifts of the early K intermediate and of several L intermediates of the bR photocycle [290], and has later allowed for the characterization of the retinal and the Schiff base chemical shifts and retinal conformations for these states [291]. Green proteorhodopsin (PR), the first functional homologue of bR to be discovered in the eubacterial domain, was discovered in the year 2001 [292]. Since then, many proteins homologous to PR have been found and are widely distributed in most of the taxonomic subdivisions of oceanic bacteria [293]. Both homology modeling and FTIR spectroscopy point to several unique features of PR. For example, the active core of PR contains polar residues, e.g., His75, which was shown to interact with proton acceptor Asp97 [294], and a few asparagines located in the cytoplasmic half of helix G, not observed in bR [295–297]. Additionally, the extracellular pair of glutamates Glu194 and Glu204, constituting the proton-releasing complex in bR [298] is replaced by hydrophobic leucines in PR, implying a different mechanism of proton ejection. Backbone and side chain chemical shift assignments have been obtained for PR in DMPC/DMPA lipids using 3D heteronuclear dipolar correlation MAS ssNMR experiments on one uniformly 15N,13C labeled sample, and two reversely 15N,13C labeled samples [108,110]. Chemical shifts have identified helical boundaries for most of the helices, a peripheral helix in the cytoplasmic E–F loop, and a short b-turn in the extracellular B–C loop (Fig. 8), consistent with the common microbial rhodopsin architecture [42,99,109, 299]. Additional assignments for mobile residues were obtained using J-coupling driven correlation spectroscopy [300]. Perdeuterated back-exchanged PR was also studied by protondetected ssNMR [301]. Like many other membrane proteins, PR has a well-protected hydrophobic core, which is inaccessible to solvent, and could not be back-exchanged. Similar effects were observed in bR [302]. While the majority of the approximately 100 resonances detected and assigned in ssNMR spectra of PR corresponded to residues in the loops and in the exposed flanks of helices, some amino acids located in the extracellular half of helix G, deep inside the lipid bilayer, could be detected as well (Fig. 8) [301]. These residues form a hydrophilic face inside the a-helical bundle, indicating the presence of a hydrophilic cavity on the extracellular side, and may in principle be involved in mediating proton release in PR.

Several sensory rhodopsins have been investigated using ssNMR techniques. Sensory rhodopsin II from N. pharaonis (SRII) is a membrane embedded photosensor that mediates a photophobic response to blue/green light. SRII changes its conformation upon illumination, and initiates conformational changes in its membrane-associated cognate transducer (HtrII). The SRII–HtrII complex extends into the cytoplasm, where it interacts with a His-kinase that phosphorylates a regulatory protein responsible for phototaxis (Fig. 7). Baldus and coworkers used MAS ssNMR in combination with reverse labeling to study SRII in native bacterial purple membranes [109]. Assignments, which were obtained for about one third of the protein residues, indicated that the secondary structure was in agreement with the X-ray structure. The use of through-bond and through-space polarization transfer mechanisms for spectral excitation and obtaining 2D correlation spectra allowed mobile and immobile fragments to be differentiated, specifically identifying three rigid loops, which could be buried in the membrane as indicated by water-edited experiments. Comparisons of the spectra collected from an unbound SRII sample, and from the SRII/HtrII complex revealed that the SRII interaction interface extends well beyond what had been previously identified from the crystal structure of the SRII/HtrII complex [303], and involves the cytoplasmic side of helix F and the E–F loop. A comparison between spectra excited using through-bond INEPT and through-space cross-polarization also revealed changes in the dynamic behavior of both SRII and HtrII upon complexation: the dynamic motions of the E–F loop of SRII became restricted in the complex, while HtrII underwent a transition from a flexible, mobile state to a rigid a-helical structure. Chemical shift perturbations in the D75N SRII mutant (D75N mutation in combination with freeze trapping stabilize the M intermediate state) upon light activation revealed a number of localized structural perturbations in the vicinity of the retinal binding pocket in helices C, E, F and G (Fig. 9), overall suggesting tilting of helix F, and movements of helices E and G in response to illumination. Anabaena Sensory Rhodopsin (ASR) from Anabaena sp. PCC7120 is another sensory rhodopsin that has been extensively investigated by ssNMR [78,99,234,249]. It has been suggested that ASR coexpresses with a soluble transducer (ASRT), a tetrameric protein of primarily b-structure (Fig. 7). One of the early proposals suggested that both ASR and ASRT are involved in the regulation of the genes responsible for the expression of light harvesting proteins in Anabaena [304]. ASR was later shown to interact with ASRT


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Fig. 9. Solid-state NMR studies of the SRII/HtrII complex. (A) Two-dimensional 13C–13C correlation spectrum of the unbound SRII receptor (black), and in complex with unlabeled transducer (green). Chemical shift changes of labeled residues indicate that they are involved in the interface. (B) Light activation of the complex. 13C–13C difference spectra between the ground and the excited state using medium mixing time (left panel, mixing time of 40 ms) and long (right panel, mixing time of 350 ms). Red and green contours represent signals that were stronger in the ground or excited state, respectively. (C) Conformational changes in the SRII upon forming a complex with HtrII and light activation. Residues shown in blue experience chemical shift perturbations upon HtrII binding. Residues shown in green show reduced mobility in the complex. Residues undergoing chemical shift changes upon light activation are shown in red, and retinal is shown in purple. Reprinted from Structure 18, M. Etzkorn, K. Seidel, L. Li, S. Martell, M. Geyer, M. Engelhard, M. Baldus, ‘‘Complex formation and light activation in membrane-embedded sensory rhodopsin II as seen by solid-state NMR spectroscopy’’, 293–300, Copyright 2010, with permission from Elsevier.

in vitro in a light-dependent manner: ASRT is released upon light activation of ASR [305], while independent solution NMR studies identified the mechanism of DNA binding by ASRT [306]. ASR reconstituted in DMPC/DMPA lipids resulted in spectra of excellent resolution [99]. Using 2D and 3D chemical shift correlation experiments on uniformly and alternately labeled samples, approximately 90% of the backbone and side chain assignments

were completed [78,99]. The excellent spectral resolution of ASR has greatly assisted in distance measurements: a number of unambiguous long-range interhelical restraints could be determined from 2D PDSD and CHHC experiments, and from 3D HBR2 experiments, and were sufficient to calculate a low-resolution template model of ASR and to complete iterative structure calculation [78]. Intermonomer PREs in combination with circular dichroism

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studies in the visible range have established that ASR forms trimers in lipids, and have provided structural restraints for the intermonomer interface [234]. A total of 211 long-range interhelical restraints including 6 intermonomer contacts, 435 medium range and 1390 short range internuclear distances, and 24 intra- and 11 intermonomer PREs were used in the structure calculation, and resulted in a family of structures with a backbone resolution of 0.6 Å (Fig. 10) [78]. Underscoring the importance of studying membrane proteins in lipids, the trimeric arrangement of ASR was found to be completely different from the dimeric structure that had been determined by protein crystallography [307]. Dipolar order parameter and transverse relaxation measurements in ASR suggested that TM helices undergo collective fluctuations on the tens of nanoseconds timescale, while the time scale of collective motions for the extracellular B–C and F–G loops approaches hundred nanoseconds [308]. Hydrogen–deuterium exchange ssNMR experiments have provided a glimpse into the structural rearrangements in the receptor upon illumination. 2D and 3D NMR spectra have shown that only a handful of amide sites, all located at the periphery of the protein, were amenable to H/D exchange in the dark [99]. The H/D exchange pattern looked quite different under illumination conditions (Fig. 10D). It was hypothesized that significant changes in the solvent accessibility could be caused by a movement of the seventh helix G in one (or more) of the intermediate states, which resulted in the formation of a hydrophilic cavity on the cytoplasmic side of the protein, allowing water to move in [249]. 4.3. Human phospholamban Human phospholamban (PLN) is a 52-residue integral membrane protein that is a reversible regulator of Ca2+ cardiac sarcoplasmic reticulum ATPase (SERCA) activity. Monomeric PLN binds SERCA and inhibits its enzymatic activity, while phosphorylation of PLN at either Ser16 or Thr17 reverses this inhibition. PLN is subdivided into four domains: the cytoplasmic region, Ia, (residues 1– 16), the loop region (residues 17–22), domain Ib (residues 23–30), and the TM region II (residues 31–52). In membranes, wild-type PLN (WT-PLN) forms pentamers, which mostly function as storage for active monomers, but were also proposed to perform ion channel activity [309]. The many different structures of PLN agree well with the a-helical structure in the transmembrane domain, but disagree in the cytoplasmic domain, which varies its conformation according to its environment, lipid composition, protein/lipid ratio, phosphorylation state and interacting partners. In particular, early solution NMR study supported the bellflower model, in which alpha-helical cytoplasmic domains are oriented along the bilayer normal [310], while later studies suggested that the PLN pentamer adopts a pinwheel model, in which the protein has an L-shape topology, with the cytoplasmic domain oriented along the bilayer surface [258]. The high-resolution structure of monomeric PLN in DPC micelles and DOPC lipids was determined by combining solution NMR restraints and orientation ssNMR restraints, obtained on a fully functional monomeric mutant, AFA-PLN with three Cys residues mutated to Phe and Ala [87]. In DPC micelles, the AFA-PLN structure contained two a-helices connected by a semiflexible loop. Orientation restraints measured on PLN in DOPC lipids revealed that the TM helix of AFA-PLN crosses the membrane at a tilt angle of 24°, and the cytoplasmic amphipathic helix lies nearly parallel to the bilayer at an angle of 102° with respect to the membrane. This L-shaped topology supported the pinwheel model, and agreed with the earlier studies using 15N OS-ssNMR on a selectively labeled PLN, and with EPR data [311–313]. A similar hybrid approach, combining solution and solid state NMR, was used to obtain the pentameric structure of WT-PLN.

Each monomer within a pentamer adopts an L-shaped topology similar to that of monomeric AFA-PLN [258]. The conformation and orientation of the cytoplasmic helix was determined from protein–detergent NOEs from solution NMR, and further validated using 15N–1H dipolar coupling constants derived from PISEMA experiments performed on the protein in the DOPC lipid bilayer. The L-shaped monomers are arranged in a pentameric pinwheel structure, as determined using intermonomer restraints derived from a solution NMR NOESY of detergent-solubilized WT-PLN, and MAS DARR experiments (Fig. 11). The pinwheel architecture was further validated by the PRE effects caused by Gd3+ in the buffer, and by EPR restraints [313]. The pore diameter in the center of the pentameric helical bundle was only 3 Å, with hydrophobic residues facing the interior on the pore, making it energetically unfavorable for ions to pass through, and ruling out the possibility of an ion channel function of the pentamer [258]. Monomeric PLN exists in an equilibrium between a dynamically relaxed R state and a more motionally restricted T state, where the relative population of states may be affected by the lipid headgroup and charge [314]. In the T-state, which is the dominant conformation in DPC micelles and DOPC lipids, the cytoplasmic domain forms an amphipathic helix, which associates with the membrane [314,315]. In contrast, the cytoplasmic domain becomes unstructured and highly dynamic in DMPC lipids, potentially representing the R-state [174]. The conformational flexibility in DMPC may further be dependent on the protein:lipid ratio, as demonstrated by 2D DARR experiments conducted by Veglia and co-workers, which suggested two conformations of the cytoplasmic domain with different mobilities [314]. Neither of these conformations was however similar to the a-helical structures of the cytoplasmic domain observed in DPC micelles and DOPC lipids [87]. Interactions with SERCA have major effects on the PLN structure. The cytoplasmic domain of AFA-PLN undergoes major conformational change upon binding SERCA in DPC micelles, switching to an extended form, probably required for interactions with the cytoplasmic domain of SERCA, and a similar transition was detected in lipids [316]. Solid state NMR on AFA- and WT-PLN in DOPC lipids shows that PLN is immobilized in the SERCA/PLN complex [315]. Upon phosphorylation of WT-PLN at S16, the cytoplasmic domain undergoes a rotation of about 20° [317]. In solution, phosphorylation disrupts the L-shaped structure of the monomeric form, shortens the cytoplasmic helix, and enhances the flexibility of the cytoplasmic domain [318]. Phosphorylation of PLN in the SERCA/PLN complex does not dissemble the complex, but loosens the contacts between PLN and SERCA [319]. 4.4. Potassium channels Among the family of potassium channels, the bacterial K+ channel KcsA has been extensively investigated as a model system for the channels function. Several factors can affect KcsA activity, e.g., K+ concentration, pH conditions and binding with channel blockers. The KcsA channel binds K+ ions through a highly conserved TVGYG sequence in the selectivity filter, which is located in the extracellular half of the protein. The selectivity filter constitutes the inactivation gate and, together with a short pore helix, connects the outer and inner membrane helices. The intracellular halves of the inner helices make up a second activation gate. The McDermott lab has carried out extensive studies of the functional dynamics and inactivation mechanisms of KcsA [75,79,165,320,321]. Wild-type, full-length KcsA was reconstituted in DOPE/DOPS liposomes, and partial spectroscopic assignments for the functionally important residues located in the selectivity filter were initially obtained using 2D and 3D heteronuclear correlation spectroscopy [321]. Two sets of chemical shifts were observed with changes in K+ concentration, with the strongest perturbation


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Fig. 10. Structure determination of ASR. (A) Aromatic–aliphatic region of a 2D PDSD 13C–13C correlation spectrum recorded at 800 MHz with a mixing time of 500 ms. Cross peaks corresponding to long-range interhelical contacts are indicated. (B) Family of ten lowest energy structures determined by MAS ssNMR (PDB ID: 2M3G). (C) Comparison of the oligomeric arrangement in the X-ray structure (left), and in the ssNMR structure (right). (D) Hydrogen/deuterium exchange of ASR [78,249]. Residues exchangeable in the dark (left) and under illumination (right) are shown in blue. Reproduced from Nature Methods 10, ‘‘Solid-state NMR spectroscopy structure determination of a lipidembedded heptahelical membrane protein’’, S. Wang, R.A. Munro, L. Shi, I. Kawamura, T. Okitsu, A. Wada, S.Y. Kim, K.H. Jung, L.S. Brown, V. Ladizhansky, 1007–1012 (2013).

being detected for residues V76 and G77, which reside in the middle of the selectivity filter. Solid state NMR titration experiments showed that KcsA has a strong potassium affinity with a dissociation constant in the range of a few lM. Low K+ concentration induces a collapsed state of KcsA, and the analysis of the effects of the E71A mutation on the chemical shift perturbation pattern led to the conclusion that the structure of the selectivity filter is similar to that of a low pH C-type inactivated state [75]. The acquisition of nearly complete assignments of the channel allowed extending information on the global structural rearrangements occurring in the channel in the low potassium state [79]. Additional chemical shift perturbations were detected in the intracellular pH gating region, and in the hinge region of the intracellular helix, altogether pointing at allosteric coupling across the membrane, and at a global conformational change which accompanies ion release. Baldus and co-workers have been studying a chimeric potassium channel protein KcsA-Kv1.3, which differs from KcsA by 11 residues, mostly located in the pore region. This modification to the sequence makes the channel a high-affinity receptor to scorpion toxins, similar to the eukaryotic voltage-gated Kv1.3 channel [72,98,322]. Interaction and the mechanisms of inhibition by two different channel blockers, 38-residue long scorpion toxin kaliotoxin (KTX) [72], and a tetraphenylporphyrin derivative (porphyrin), were studied. Chemical shift mapping identified the interaction interface of the KTX-KcsA-Kv1.3 complex, which included residues on the extracellular surface of the channel

(Fig. 12), while the conformation of the selectivity filter resembled that of the closed-conductive state. In contrast, porphyrin was found to bind in the middle of the pore region of KcsA-Kv1.3, leading to the collapse of the selectivity filter, and showing an inhibition mechanism similar to the pH dependent gating of KcsA [323]. The secondary structure of KcsA-Kv1.3 in asolectin liposomes derived from chemical shift assignments mostly agreed with the crystal structure of KcsA, obtained at high potassium concentration [324], with the exception of two N- and C-terminal amphipathic helices, and a longer outer transmembrane helix [98]. Chemical shift data and 2D NHHC and CHHC experiments carried out under a high potassium concentration of 50 mM, showed that the conformation of the selectivity filter reproduces that of the conductive state, which is an active conformation for K+ transport [325]. The structural mechanisms of the pH dependence of KcsA gating have also been investigated. Comparison of 2D PDSD spectra of KcsA-Kv1.3 at different pH values and at a high potassium concentration showed that the channel gating involves structural changes in the selectivity filter, pore helix and inner transmembrane helix [323,325]. A decrease in the pH leads to the collapse of the selectivity filter, and to bending of the inner TM helix in the intracellular half of the channel, overall resulting in the opening of the activation gate [323]. Based on this observation, an allosterically coupled gating mechanism was suggested, in which the collapse of the selectivity filter opens the activation gate, and the conducting conformation of the selectivity filter closes the activation gate (Fig. 12) [323].

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Fig. 11. Pentameric structure of phospholamban. (A) Two-dimensional 13C–13C correlation spectrum collected with a DARR mixing time of 200 ms on a 1:1 mixture of uniformly 13C-Leu and uniformly 13C-Ile labeled PLN. Intraresidue and interprotomer cross-peaks are shown in black and blue, respectively. (B) Overlay of 2D 1H–15N PISEMA spectra of selectively labeled PLN in oriented DOPC/DOPE lipid bilayers. (C) CSA as a function of residue number. (D) Dipolar couplings as a function of residue number. Reproduced from Proc. Natl. Acad. Sci. USA 108, R. Verardi, L. Shi, N.J. Traaseth, N. Walsh, G. Veglia, ‘‘Structural topology of phospholamban pentamer in lipid bilayers by a hybrid solution and solid-state NMR method’’, 108 (2011) 9101–9106, The National Academy of Sciences of the USA, 2011. (E) Pentameric structure of PLN obtained using solution and solid-state NMR restraints (PDB ID: 2KYV).

Fig. 12. (A–D) Interaction between KscA-Kv1.3 and kaliotoxin. (A) 2D 13C–13C correlation spectrum of an unbound KcsA-Kv1.3 channel (red), and bound to unlabeled KTX (green). (B) and (C) highlight specific regions with affected residues. (D) Residues of the channel perturbed by KTX binding are shown in red, and unperturbed residues are shown in blue. Reprinted by permission from Macmillan Publishers Ltd.: Nature 440, ‘‘Toxin-induced conformational changes in a potassium channel revealed by solid-state NMR’’, A. Lange, K. Giller, S. Hornig, M.F. Martin-Eauclaire, O. Pongs, S. Becker, M. Baldus, 959–962, Copyright 2006. (E–F) Effects of potassium concentration and pH on the KscA-Kv1.3 activation. (E) Left panel: comparison of 2D 13C–13C spectra at pH 7.5 (black) and pH 4.0 (blue) at low [K+]. Perturbed residues T74, T75, V76 are part of the selectivity filter, and T101 is in the gating hinge. Right panel: comparison of 2D 13C–13C spectra at pH 7.5 (black) and pH 4.0 (red) at 50 mM [K+]. (F) Cartoon representation of KcsA–Kv1.3 in a lipid bilayer. The selectivity filter acts as an inactivation gate and resides in either conductive/open or collapsed/closed states. The activation gate is located in the TM2 helix of each subunit and gate opening is associated with a bent helix. Reprinted from ‘‘Coupling of activation and inactivation gate in a K+-channel: potassium and ligand sensitivity’’, C. Ader, R. Schneider, S. Hornig, P. Velisetty, V. Vardanyan, K. Giller, I. Ohmert, S. Becker, O. Pongs, M. Baldus, EMBO J., 28, 2825–2834. Copyright 2009, John Wiley & Sons, Inc.

In contrast, a high K+ concentration of 50 mM stabilized the closed conductive conformation of the selectivity filter regardless of pH [323]. Analysis of the chemical shifts of the side chain

carboxyls of E71, E118 and E120 revealed that they were deprotonated at acidic pH, and protonated in the absence of K+ [323]. These residues are either involved in the hydrogen bonding network


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Fig. 13. Structure and topology of DsbB. (A) 2D 13C–13C correlation spectrum excited by proton spin diffusion from lipid acyl chains to DsbB. (B) 2D 13C–13C correlation spectrum excited by proton spin diffusion from water to DsbB. (C) Structural model of DsbB calculated using joint X-ray and ssNMR restraints, and refined in the bilayer using molecular dynamics. Residues shown in orange correlate with lipids and show up in the spectrum in (A). Residues shown in blue correlate with water and show up in (B). Water and lipid molecules are shown in cyan and gray, respectively. Reprinted from Journal of Molecular Biology, 425, M. Tang, A.E. Nesbitt, L.J. Sperling, D.A. Berthold, C.D. Schwieters, R.B. Gennis, C.M. Rienstra, ‘‘Structure of the disulfide bond generating membrane protein DsbB in the lipid bilayer’’ 425, 1670–1682, Copyright 2013, with permission from Elsevier.

stabilizing the selectivity filter, or are located in the inactivation gate region and play an important role in both activation and inactivation gating mechanisms [323]. 4.5. DsbB–DsbA disulfide bond formation complex Dsb enzymes located in the periplasmic space of E. coli catalyze the formation of disulfide bonds in E. coli envelope proteins. The system consists of a soluble enzyme DsbA, which oxidizes the free Cys residues of targeted proteins, and a cytoplasmic membrane protein DsbB, which transmits electrons from DsbA to ubiquinone (UQ). The structure of DsbB and the DsbA–DsbB complex and the mechanistic details of the complex function have been investigated by MAS ssNMR [76,97,118,259,326]. The C41S DsbB mutant, which mimicks a transient intermediate state in the reaction pathway, was one of the first transmembrane proteins for which nearly complete assignments were obtained using a single uniformly 13 15 C, N-labeled sample [97]. The 3D heteronuclear correlation experiments were carried out on a precipitated protein, with only endogenous E. coli lipids present, ensuring that large amount of sample (1 lmol) could be packed into an NMR rotor. The high-resolution ssNMR/X-ray structure of precipitated DsbB was determined using a combined set of more than 600 NMR distance restraints obtained from sparsely labeled samples grown on either 2-13C glycerol or 1,3-13C glycerol, torsional TALOS restraints, and X-ray reflections [76]. This combined structure refinement approach had been previously demonstrated by the same group on the DsbA–DsbB complex to result in an improved quality of structure when compared to X-ray constraints alone [259]. To account for the effect of the bilayer, the structure was further refined using molecular dynamics of the protein in POPE lipids. To confirm the topology of the protein, additional water- and lipid-edited experiments were recorded on the DsbA–DsbB complex reconstituted in the phospholipid bilayer, and revealed residues in the vicinity of the protein–solvent and protein–lipid interfaces (Fig. 13). The C41S mutant of DsbB was also used to investigate the charge-transfer complex between C44 of DsbB and UQ-8 [327]. The formation of the complex was detected by both the unique chemical shifts of I45 and by a maximum absorption at 500 nm in UV/Vis spectroscopy [327]. Full isoprenoid side chain assignments and partial assignments of the quinone headgroup of UQ-8 in the Cys-UQ-8 complex were obtained using double-quantum spectroscopy, guided by solution NMR assignments. A comparison

of NMR spectra at high pH 8 where the Cys-UQ-8 complex is stable, and at pH 5.5, at which the complex dissociates, revealed the different protonation states of the thiol group of C44. In the deprotonated state at high pH, the negatively charged sulfur atom transfers an electron to the carbonyl of UQ, thus stabilizing the complex. In contrast, at lower pH, the partial negative charge of the thiolate of C44 results in a reduced electron density of the quinone group, thus weakening the thiol-UQ-8 interactions. 4.6. G-protein coupled receptors G-protein coupled receptors are seven transmembrane helical receptors which respond to a large array of extracellular stimuli and transmit information into the cell. With nearly 1000 members, GPCRs represent the largest membrane protein family in the human genome. They are associated with many diseases, and many of them represent drug targets. In the past five years there has been remarkable progress in the structure determination of GPCRs by X-ray crystallography [8–18], as well as in the characterization of GPCRs by solution NMR [45–47,328]. As lipids may play various roles in GPCR function, it is important to study these proteins in the lipid environment, and they are attractive targets for solid-state NMR spectroscopy. Owing to difficulties with expression and isotopic labeling of GPCRs, the majority of these studies have focused on samples with selective labels, or on ligands bound to GPCRs [329–331]. Visual rhodopsins are visual pigments in the photoreceptor cells in the retina of the eye. They are responsible for the first event in the perception of light. Rhodopsin binds 11-cis retinal as its chromophore and is very sensitive to light, enabling vision in low light conditions. Absorption of light causes isomerization of the retinal, and triggers a light cycle which includes a number of intermediate states, of which Meta II is the active state and Meta I is its precursor. Using different site-specific labeled rhodopsin samples expressed in the HK293b cell line, Smith and coworkers investigated the mechanism of activation in bovine rhodopsin [140– 146]. The Meta II state was trapped by freezing rhodopsin solubilized in detergent after illumination, and the formation of this intermediate state was confirmed by changes in the retinal chemical shift [144]. 2D DARR experiments on the rhodopsin regenerated with 13C-labeled retinal, suggested that the C20 methyl group of the retinal rotates by about 90°, and the b-ionone ring of the retinal moves 4–5 Å closer to helix 5 [142,144], which was consistent with the counterion switch model proposed for

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Fig. 14. 2H NMR relaxation studies of activation mechanism of rhodopsin. (A) Order parameters (SC3), pre-exponential factors (k0) for three-fold axial jumps or D0 for continuous diffusion, and activation energies Ea for C5-Me, C9-Me, and C13-Me methyl groups of retinal in the dark, Meta I and Meta II states. (B–D) Proposed activation mechanism for rhodopsin. Isomerization of retinal displaces the extracellular loop 2 toward the extracellular side (e side). Collective fluctuations of helices H5 and H6 expose transducin recognition sites on the cytoplasmic side (c side). Reprinted by permission from Macmillan Publishers Ltd.: Nature Structural Molecular Biology 18, ‘‘Retinal dynamics underlie its switch from inverse agonist to agonist during rhodopsin activation’’, A.V. Struts, G.F. Salgado, K. Martinez-Mayorga, M.F. Brown, 392–394, Copyright 2011.

Fig. 15. Structure determination of CXCR1. (A) Topology of CXCR1. Cysteines are shown in gold, disulfide bonds are dashed lines. (B) Representative strip plots from the 3D NCACX experiment for residues 31–35 in the N-terminus (red), residues 78–82 of the transmembrane helix 2 (blue), and residues 175–179 of the second extracellular loop. (C) 13C detected 1H–15N separated local field spectra. (D) 1H–15N dipolar couplings as a function of residue number. Sinusoidal fits in cyan are shown for seven TM helices and the C-terminal helix 8 (H8). (E) Ensemble of 10 lowest energy structures. (F–G) Experimental vs back-calculated dipolar couplings before (F) and after (G) refinement against the experimental data. Reprinted by permission from Macmillan Publishers Ltd.: Nature 491, ‘‘Structure of the chemokine receptor CXCR1 in phospholipid bilayers’’, S.H. Park, B.B. Das, F. Casagrande, Y. Tian, H.J. Nothnagel, M. Chu, H. Kiefer, K. Maier, A.A. De Angelis, F.M. Marassi, S.J. Opella, 779–783, Copyright 2012.


rhodopsin activation [332]. 1D and 2D ssNMR on differently isotopically labeled samples showed that this movement induced a number of structural rearrangements within the protein, including the displacement of the second extracellular loop [146], the displacement and rotation of helix 5 [144,146], and an outward rotation of helix 6 [144]. Solid state NMR data also suggested that the formation of the Meta II state breaks the characteristic ionic lock structure, composed of charged residues on helix 3 (E134 and R135) and helix 6 (E247), and results in an interhelical contact between R135 and M257 of helix 6 [140]. Critical factors stabilizing the Meta II active conformation have also been identified from ssNMR measurements [140]. For example, the highly conserved Y223 was found to face the interior of the lipid bilayer in the dark state, but upon light activation changed its position towards the protein core. To gain insights into the functional dynamics of the activation events in rhodopsin, Brown and co-workers have conducted 2H relaxation measurements [333,334]. Three methyl groups of the retinal were deuterated at positions C5, C9, and C13 and their relaxation rates were measured in the dark, Meta I, and Meta II states. Relaxation rates were analyzed in terms of a three-site jump model, and revealed major differences in the pre-exponential factors and rotation activation barrier energies (Fig. 14), reflecting differential mobility of the retinal in the dark and in the intermediate states. These data, coupled with molecular dynamics simulations, have allowed the researchers to propose a multiscale activation mechanism, where retinal isomerization initiates the collective motions of helices 5 and 6 on the microsecond-to-millisecond time scale with multiple activated substrates in the Meta I–Meta II equilibrium (Fig. 14). In the past few years preceding this review, several groups have attempted to express isotopically labeled GPCRs in E. coli for solidstate NMR. Gawrisch, Yeliseev and co-workers, and Huster and co-workers have published expression protocols for the Human Cannabinoid Type 2 Receptor, and for the Neuropeptide Y2 Receptor [122,123,335]. Although good yields were obtained for both proteins, and the first multidimensional correlation spectra were recorded, the observed resolution was visually worse than that published for other proteins of comparable size, notably 7TM bacterial rhodopsins, thus making site-specific assignments challenging. The chemokine CXCR1 receptor is the only example of a successful ssNMR structure determination of a GPCR. Opella and coworkers over-expressed CXCR1 in E. coli, refolded and reconstituted it in DMPC liposomes [82,125]. Above the phase transition temperature of DMPC, CXCR1 forms a rigid helical bundle with flexible termini which undergoes fast rotational diffusion in a phospholipid bilayer [336]. This results in partial averaging of anisotropic dipolar interactions, and allows rotationally aligned (RA) ssNMR to be utilized for the determination of the backbone structure of the protein. RA ssNMR combines the advantages of magic angle spinning NMR and oriented-sample solid-state NMR in that it allows for both the isotropic chemical shifts and motionally averaged 15N–1H and 13 C–1H dipolar couplings to be measured simultaneously, also taking advantage of higher sensitivity of carbon detection (as opposed to 15N detection as is typically used in OS NMR) [81]. In CXCR1, the dipolar wave pattern identified seven TM helices, some of which contained kink distortions, and an amphipathic helix positioned along the bilayer (Fig. 15). Using the visual rhodopsin structure as a template, 1H–15N and 1H–13C dipolar couplings determined the backbone 3D structure of CXCR1, with an overall topology consistent with that of the common GPCRs. For example, a helical kink detected in the second helix of CXCR1 is also found in the crystal structure of another chemokine receptor CXCR4. Likewise, the b-hairpin structure detected in CXCR1 is found in CXCR4 as well as in other GPCRs. The CXCR1 structure appears to be exceptionally rigid. As evident from the examination of the family of structures in Fig. 15E,

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the loop conformations are well defined, as reflected by relatively low RMSD for these regions. The reduced mobility of the loops is also consistent with the fact that nearly complete spectroscopic assignments could be obtained for these regions. In contrast, loop regions of GPCRs are less ordered/floppier in crystal structures, and often need to be modified for successful crystallization. Similarly, the majority of ssNMR studies performed on membrane proteins either reconstituted in lipid bilayers or prepared in other forms indicate increased mobility of loop regions. Extending the library of GPCR structures determined in the lipid environment would help determine whether the apparent rigidity of the loop regions of CXCR1 is an intrinsic property of this particular receptor, or is related to the state of the protein in lipids. 5. Concluding remarks and perspectives In this review we have briefly discussed the recent progress in solid-state NMR approaches for the characterization of membrane proteins, and have given a few examples where this technique has provided especially important structural and functional insights, often undetectable by other conventional high-resolution structural methods. These studies demonstrate that solid-state NMR can be successfully used to investigate membrane proteins in a native-like lipid environment under nearly physiological conditions. Well-resolved spectra obtained for different classes of membrane proteins indicate that the sample preparation for membrane proteins appears to be more straightforward than for X-ray crystallography or solution NMR, as it relies on the native property of membrane proteins to favor lipid environments. Methods for both oriented sample and magic angle spinning NMR are available now. Sensitivity, which is already sufficient for many applications, will continue to improve through the development of proton detection methods [337]. Additionally, Dynamic Nuclear Polarization has great potential for providing large signal enhancements [290,291,338], although its application for a complete site-specific structural analysis is limited at present by the line broadening under low temperature conditions. The development of fast magic angle spinning and perdeuteration allows the intrinsic problem of short coherence life times in solid-state NMR to be overcome, and holds great promise for studies of protein dynamics [308,339– 341]. Thus, these developments suggest that solid-state NMR contributions to membrane protein structural biology will expand in the next few years and will complement other powerful highresolution methods. Acknowledgements This work was supported by NSERC Discovery grant to V.L., and start-up funds from Peking University to S.W. V.L. holds Canada Research Chair Tier II in Biophysics. We are grateful to Ms. Meaghan Ward for carefully reading the manuscript and providing useful suggestions. We thank Drs. M. Baldus (Utrecht University), B.-J. van Rossum (Leibniz-Institut für Molekulare Pharmakologie), C.M. Rienstra (University of Illinois at Urbana Champaign), and M. Tang (City University of New York) for providing us with high-resolution figures for this review. References [1] E. Wallin, G. von Heijne, Genome-wide analysis of integral membrane proteins from eubacterial, archaean, and eukaryotic organisms, Protein Sci. 7 (1998) 1029–1038. [2] R.M. Stroud, New tools in membrane protein determination, F1000 Biol. Rep. 3 (2011) 8. [3] R.M. Bill, P.J. Henderson, S. Iwata, E.R. Kunji, H. Michel, R. Neutze, S. Newstead, B. Poolman, C.G. Tate, H. Vogel, Overcoming barriers to membrane protein structure determination, Nat. Biotechnol. 29 (2011) 335–340.

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Glossary of abbreviations ASR: Anabaena Sensory Rhodopsin ASRT: Anabaena Sensory Rhodopsin transducer Amt: amantadine bR: bacteriorhodopsin CXCR1: chemokine (C-X-C motif) receptor 1 DsbB: Disulfide bond formation protein B DARR: dipolar assisted rotational resonance DMPA: 1,2-dimyristoyl-sn-glycero-3-phosphate DMPC: 1,2-dimyristoyl-sn-glycero-3-phosphocholine DMPG: 1,2-Dimyristoyl-sn-Glycero-3-phospho glycerol DOPS: 1,2-dioleoyl-sn-glycero-3-phospho-L-serine DOPE: 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine DPC: n-dodecylphosphocholine EDTA: ethylenediaminetetraacetic acid GPCR: G-protein coupled receptor HBR2: homogeneously broadened rotational resonance POPS: 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-L-serine POPE: 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine PA: phosphatidic acids PS: L-a-phosphatidylserine M2TM: M2 transmembrane domain M2CD: conduction domain MAS: Magic Angle Spinning MTSL: (S-(2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrol-3-yl)methyl methanesulfonothioate NMR: nuclear magnetic resonance OS NMR: oriented sample NMR PCS: pseudocontact shift PDSD: proton driven spin diffusion PISEMA: Polarization Inversion Spin Exchange at the Magic Angle PLN: phospholamban PR: proteorhodopsin PRE: paramagnetic relaxation enhancement R2: rotational resonance RA: rotationally aligned REDOR: Rotational Echo DOuble Resonance SERCA: sarcoendoplasmic reticulum calcium transport ATPase SDS: sodium dodecyl sulfate SRII: sensory rhodopsin II ssNMR: solid-state NMR TEDOR: Transferred Echo DOuble Resonance TM: transmembrane 7TM: seven-helical WT: wild type YadA: Yersinia adhesin A

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