Abstract
Based on continuous methodical advances and developments, solid-state NMR spectroscopy has become a powerful tool for the investigation of various materials, including polymers, glasses, zeolites, fullerenes, and many others. During the past decade, solid-state NMR spectroscopy also found increasing interest for the study of biomolecules. For example, membrane proteins reconstituted into lipid environments such as bilayers or vesicles, protein aggregates such as amyloid fibrils, as well as carbohydrates can now be studied by solid-state NMR spectroscopy. This review briefly introduces the principles of solid-state NMR spectroscopy and highlights novel methodical trends. Selected applications demonstrate the possibilities of solid-state NMR spectroscopy as a valuable bioanalytical tool.
Similar content being viewed by others
Avoid common mistakes on your manuscript.
Introduction
Bioanalytics deals with the detection and characterization of proteins, nucleic acids, carbohydrates, and lipids. Liquid-state nuclear magnetic resonance (NMR) spectroscopy has become one of the most successful techniques for investigations of the structure and dynamics of biomolecules in solution [1, 2]. During the past decade, solid-state NMR spectroscopy found an increasing number of biological applications, for example, the study of membrane proteins in lipid environments (see Fig. 1), protein aggregates such as amyloid fibrils, as well as crystalline proteins or protein precipitates [3–22]. Biomineralization phenomena can be studied by solid-state NMR spectroscopy as well [23], including investigations on integral cells [24].
In special cases, conventional one-dimensional NMR spectroscopy allows the detection and characterization of certain biomolecules. A recent example is shown in Fig. 2. Unexpectedly, chitin-based scaffolds could be identified for the first time in the skeleton of the marine demosponge Ianthella basta by solid-state 13C NMR spectroscopy [25]. Similar scaffold structures were also observed and characterized by solid-state NMR spectroscopy in diatom cell walls [26]. The biopolymer chitin exhibits a rather simple and well-known 13C NMR spectrum; hence, the identification of the spectral fingerprint can be solely based on one-dimensional NMR spectroscopy. In most cases, however, multidimensional NMR techniques must be applied in analogy to liquid-state NMR spectroscopy to resolve the various signals and to extract structural information from the spectra. The aim of this review is to provide a brief introduction to the principles of solid-state NMR spectroscopy and to highlight emerging trends which may further enhance the bioanalytical potential of solid-state NMR spectroscopy.
Solid-state NMR—a short introduction
The first successful NMR experiments on condensed matter were performed in 1945 [27, 28]. The famous Pake doublet in solids, i.e., the line shape resulting from the magnetic dipole–dipole interaction within isolated spin pairs, was discovered in 1948 for the 1H NMR signal of water molecules in hydrated crystals such as gypsum [29]. Apart from the homonuclear and heteronuclear magnetic dipole–dipole interactions between neighboring nuclei, the chemical shift anisotropy and—for nuclei with I > 1/2—the electric quadrupole interaction are the dominating line-broadening interactions in solids [30]. The measurement of chemical shifts in solids, therefore, mandatorily requires the application of line-narrowing techniques to suppress the aforementioned line-broadening interactions as much as possible. This can be accomplished by mechanical spinning of the sample [31] around an axis tilted by the magic angle ϑ m = 54.7° with respect to the external magnetic field, B 0, a technique which is denoted magic-angle spinning (MAS) or magic-angle sample spinning. Another approach for the suppression of homonuclear magnetic dipole–dipole interactions (homonuclear decoupling) is based on the irradiation of special pulse sequences [32, 33] such as the Waugh–Huber–Haeberlen (WAHUHA) sequence. Combined with MAS, combined rotation and multiple-pulse spectroscopy (CRAMPS) experiments result in an excellent spectral resolution for strongly coupled homonuclear spin systems such as 1H and 19F even if the available sample spinning rates are rather low (see Fig. 3) [33–35]. Especially in multidimensional NMR experiments, frequency-switched Lee–Goldburg (FSLG) or phase-modulated Lee–Goldburg (PMLG) decoupling or decoupling using mind-boggling optimization (DUMBO) is nowadays usually applied for homonuclear decoupling [36–38]. These sequences are designed such that the homonuclear magnetic dipole–dipole interaction is averaged out as efficiently as possible. The combination of MAS and multiple pulse sequences is necessary if the spin systems exhibit strong homogeneous interactions such as homonuclear magnetic dipole–dipole interactions among multiple spins. The suppression of these interactions would require extremely high sample spinning rates even beyond 100 kHz, which are not yet available (see later). However, the combination of MAS and the described pulse sequences results in superior resolution, as is shown in Fig. 3.
Solid-state NMR measurements of nuclei S with low gyromagnetic ratio γS such as 13C, 15 N, and 29Si using direct excitation are often extremely time-consuming (if not impossible). Their intrinsically low sensitivity is caused by the low spin polarization and sometimes low natural abundance in combination with rather long longitudinal relaxation times T 1 determining the repetition time between two subsequent scans (see later). Therefore, solid-state NMR spectra of the aforementioned nuclei are commonly acquired by cross-polarization (CP) experiments [39, 40]. The CP experiment is a typical polarization transfer experiment. Nuclei I of high magnetogyric ratio γI such as 1H and 19F typically serve as the source of magnetization, which is transferred to neighboring nuclei S. The physical basis of this polarization transfer is the heteronuclear magnetic dipole–dipole interaction between I and S. This experiment can result in a maximum signal enhancement given by γI/γS. For a 13C{1H} CP experiment (S is 13C and I is 1H), this maximum enhancement factor amounts to approximately 4. Since 1H nuclei often exhibit a relatively short T 1 compared with the S nuclei, the effective signal-to-noise improvement factor within a given measurement time is usually much larger than γI/γS (see later). Ramped or adiabatic-passage CP experiments are nowadays often preferred [41–43]. Heteronuclear 1H decoupling sequences such as two-pulse phase-modulated (TPPM) and small-phase incremental alternation (SPINAL) decoupling [44, 45] are commonly applied during signal acquisition to suppress the influence of the strongly coupled 1H nuclei upon the line width of the S signals (13C, 15 N, 29Si, 31P, etc.). Figure 4 demonstrates the application of the described techniques for the measurement of highly resolved solid-state 31P NMR spectra of crystalline O-phospho-l-tyrosine. Note the shape of the 1H-decoupled 31P NMR signal measured without MAS which is mainly determined by the chemical shift anisotropy.
Fast MAS results in the removal of the influence of line-broadening interactions from the spectra. Especially the distance-dependent dipolar couplings are, however, a valuable source of structural information. Reintroduction of dipolar couplings allows the determination of internuclear distances. One frequently used technique is the rotational echo double resonance (REDOR) experiment [47] to selectively determine heteronuclear dipolar couplings. Rotational resonance experiments [48] allow the reintroduction of homonuclear dipolar couplings.
The application of multidimensional liquid-state NMR techniques [49] to proteins and other biomolecules in solution was a major breakthrough in biological NMR spectroscopy [1]. Multidimensional NMR spectroscopic techniques are also routinely used in solid-state NMR spectroscopy. Figure 5 shows a 1H-31P heteronuclear correlation (HETCOR) spectrum of the crystalline amino acid O-phospho-l-serine without and with homonuclear FSLG decoupling during the evolution time t 1. The spectra were measured under MAS at a sample spinning rate ν r of 12 kHz. Ramped 1H-31P CP was used for polarization transfer. Note the considerable improvement of resolution in the indirect spectral dimension (1H spectrum) obtained by the application of homonuclear FSLG decoupling during the evolution time. The pulse scheme used for the FSLG decoupled experiment is given at top of the figure. This type of spectroscopy allows the detection of through-space correlations, dipolar couplings between different nuclei (here 1H and 31P). Through-bond correlations, i.e., J couplings, can be detected by a solid-state NMR experiment such as the incredible natural abundance double quantum transfer experiment (INADEQUATE) [50–52]) or total through-bond correlation spectroscopy (TOBSY) [53]. Furthermore, insensitive nuclei enhanced by polarization transfer (INEPT) experiments—originally designed for heteronuclear liquid-state NMR spectroscopy [2]—are also feasible in solid-state NMR spectroscopy [54], especially under efficient homonuclear 1H decoupling and at high sample spinning rates for biological samples [55, 56].
In addition to the structural information described, solid-state NMR spectroscopy also allows the determination of dynamics such as intramolecular motions within the biomolecules. One possibility is the analysis of relaxation data, i.e., the study of the influence of thermal motions upon the longitudinal and transverse relaxation times T 1 and T 2, respectively. This could recently be applied to investigate the mobility of chondroitin sulfate in articular and artificial cartilage [57]. It should be noted that the presence of thermal motions influences the shape and line width of MAS NMR signals in a characteristic manner depending on the correlation time, τ C, of the motional processes (see “Spectral resolution”). Elaborate techniques have been developed for the study of proteins: For example, mobile protein segments can be detected using the through-bond correlation schemes introduced by Andronesi et al. [5]. Conformational dynamics in proteins with correlation times between milliseconds and seconds become detectable in dipolar centerband-only detection of exchange (CODEX) NMR experiments [58, 59] (see Fig. 6).
Sensitivity and spectral resolution are the crucial parameters which determine and limit the applicability of any spectroscopic technique. Often, the available amounts of biological samples are rather limited—in particular if isotope labeling is necessary. The detection of very small amounts of sample is, therefore, particularly important for bioanalytical applications. Furthermore, the complex biomolecules often exhibit numerous signals, resulting in resolution problems. The following sections are, therefore, devoted to the discussion of sensitivity and resolution especially with respect to novel biological applications of solid-state NMR spectroscopy.
Improvements in sensitivity
The signal-to-noise ratio obtained in an NMR experiment depends on the macroscopic magnetization of the sample, which is determined by the spin polarization P. The spin polarization is given by
for spin-1/2 nuclei. Here, N +1/2 and N -1/2 are the populations of the two Zeeman energy levels corresponding to magnetic spin quantum numbers +1/2 and -1/2, respectively. These populations are given by the Boltzmann distribution in thermal equilibrium. For spin-1/2 nuclei, one can write within the high-temperature approximation, which is valid at room temperature and the currently available magnetic fields,
ħ is the Planck constant divided by 2π, k denotes the Boltzmann constant, and T is the absolute temperature. Note that the signal-to-noise ratio also depends on various other parameters, for example, the coil filling factor, the sample volume, the noise figure of the preamplifier, and the temperature of the detection coil [60, 61]. Furthermore, the signal-to-noise ratio increases with the square root of the number of scans which are added to acquire the free-induction decay. The repetition time between two subsequent scans is limited by the longitudinal relaxation time. After a π/2 excitation pulse, polarization buildup times of about 5 times T 1 are commonly allowed before the next scan. Consequently, the signal-to-noise ratio depends on the square root of the measurement time.
As can be seen from Eq. 2, the spin polarization and hence the signal-to-noise ratio can be enhanced by increasing the magnetic field B 0. Over the past few decades, the field strength of commercial spectrometers has continuously increased. Nowadays, superconducting magnets up to approximately 23 T are available. Pulsed magnets with maximum fields up to 56 T could be developed and used for the detection of 1H NMR spectra [62]. However, the room-temperature equilibrium spin polarization P of 1H nuclei amounts to only 0.02% even at 56 T. This is the physical reason for the intrinsically low sensitivity of NMR spectroscopy. Therefore, sensitivity improvement remains one of the major methodical challenges in NMR spectroscopy [63, 64].
The spin polarization can be enhanced by sample cooling in special cases. This, however, requires very low temperatures, which cannot always be applied, especially in the case of biological samples. Furthermore, MAS at temperatures below approximately 100 K is technically difficult. The creation of nonequilibrium spin polarizations by hyperpolarization experiments is, therefore, an important field of methodical development.
Hyperpolarization methods
We have already described the advantages of the CP experiment. However, the maximum polarization gain in this experiment is limited by γI/γS. Other techniques are capable of delivering considerably higher spin polarizations. Promising hyperpolarization approaches which have already been exploited for solids or solid surfaces are dynamic nuclear polarization (DNP), chemically induced nuclear polarization (CIDNP)/photochemically induced nuclear polarization (photo-CIDNP), and spin-exchange optical pumping (SEOP).
DNP relies on the spin-polarization transfer from electrons to nuclei [65] (see also Fig. 7). In the case of diamagnetic samples, the samples have to be doped with appropriate paramagnetic agents [66]. The paramagnetic substances can be brought into contact with the molecules of interest, for example, by freezing glass-forming solutions containing paramagnetic solutes as well as the sample molecules. Spin diffusion among 1H nuclei can result in the efficient distribution of the DNP-enhanced nuclear spin polarization throughout the samples. The past few years have seen considerable methodical progress in DNP experiments [67–69] which made it possible to combine DNP with MAS. These methodical advances are currently paving the way to a variety of novel bioanalytical applications [70–72].
The CIDNP effect was discovered in 1967 independently by two groups [73–75]. It occurs as a result of radical ion reactions and is often explained by the radical pair mechanism. The electron-spin-dependent recombination probability of radical pairs is influenced by the nuclear spin, i.e., the hyperfine coupling between electrons and nuclear spins. Therefore, reactions involving radical pairs can result in strongly enhanced nuclear spin polarizations; for details, see the recent review by Bargon [76]. Photo-CIDNP was discovered shortly after the first CIDNP experiments had been performed [77, 78]. Since then, photo-CIDNP has become an important tool for photobiochemical investigations. Photo-CIDNP could also be combined with MAS [79] to study photobiochemical processes by solid-state NMR spectroscopic techniques [80–83].
SEOP is an experimental approach which can be applied to hyperpolarize noble gases such as 129Xe (see Fig. 8 as well as [84–86]). Optical pumping of alkali atoms, usually Rb, results in highly polarized electron spin systems. Electron spin polarization is then transferred to the nuclear spins by the formation of short-lived Rb–Xe van der Waals complexes or Rb–Xe collisions via hyperfine coupling. These processes lead to a 129Xe spin polarization 4 to 5 orders of magnitude higher than the equilibrium spin polarization in commonly used spectrometers. 129Xe is an important probe atom which was originally introduced by Ito and Fraissard [87] in surface NMR spectroscopy. Hyperpolarized xenon was then suggested to enhance the NMR of solid surfaces [88]. In addition, xenon is also frequently used in biological NMR spectroscopy [89–97] as well as for imaging purposes [98, 99]. SEOP, therefore, offers a variety of solid- and liquid-state NMR experiments (for reviews, see [96, 97]) even under MAS [100], including spin-polarization-transfer techniques such as the spin-polarization-induced nuclear Overhauser effect (SPINOE) [101] and spin-polarization-induced enhancement by Hartmann–Hahn dipolar recoupling (SPIDER) [102]. Biosensor applications of xenon have recently found special interest in biological liquid-state NMR spectroscopy as well as in imaging [103–105].
Enhanced spin polarizations can also be produced by hydrogenation reactions using parahydrogen [106, 107]. This parahydrogen-induced polarization (PHIP) effect can be created either in high field (parahydrogen and synthesis allow dramatically enhanced nuclear alignment, PASADENA [106]) or in low field (adiabatic longitudinal transport after dissociation engenders net alignment, ALTADENA [108]). PHIP is increasingly being exploited in liquid-state NMR spectroscopy [109], for example, in spin-polarization-transfer experiments [110, 111]. The PASADENA approach could be applied to study H2 chemisorption on ZnO [112]. Recently, Adams et al. [113] developed an approach for the reversible interaction of parahydrogen with organic substrates mediated by metal complexes. This approach results in signal enhancement factors up to approximately 800 in 1H, 13C, and 15 N NMR and can be used in magnetic resonance imaging as well. Finally, it should be mentioned that hyperpolarized spin systems provide the basis for extremely fast 2D experiments, so-called ultrafast 2D NMR spectroscopy [114]. In contrast to conventional 2D NMR spectroscopy, such techniques allow the detection of 2D spectra within a single scan or only a few scans.
T1 shortening and related approaches
As already mentioned, the signal-to-noise ratio obtained in an experiment after a certain time increases strongly if T 1 is short. T 1 shortening is, therefore, another promising approach for sensitivity enhancement. The addition of Cu(II)Na2EDTA as a chelated paramagnetic relaxation agent results in about 2 orders of magnitude shorter T 1 values for protein microcrystals [115]. Linser et al. [116] combined this approach with partial deuteration of the samples by the recrystallization of a protein in 9:1 D2O/H2O solution in the presence of the relaxation agent. Paramagnetic doping of solid proteins such as amyloid fibers and ubiquitin was combined with very fast MAS and low-power radio-frequency pulse sequences in the paramagnetic-relaxation-assisted condensed data collection (PACC) experiment [117]. This experiment results in a reduction of T 1 by orders of magnitude and allows the detection of 2D solid-state NMR spectra for protein concentrations in the nanomolar range.
Another time-saving solid-state NMR approach is the relaxation enhancement by a lower temperature of adjacent spins (RELOAD) experiment [118]. This experiment is based on the idea that selective excitation of spins such as 13C which are strongly coupled to 1H nuclei is followed by enhanced relaxation due to 1H-driven spin diffusion from nonexcited spins, thus resulting in faster magnetization recovery.
Microcoils
The sensitivity of an NMR experiment on a sample of given volume depends on the coil filling factor, i.e., the ratio between the coil volume and the sample volume. If the available sample volume is rather small, the use of microcoils results in a significant improvement in the signal-to-noise ratio. The need to study low sample volumes especially in biological NMR spectroscopy has stimulated numerous attempts to develop appropriate microcoil systems also for solid-state NMR spectroscopy [119–124]. Solenoid microcoil systems have been developed for wide-line solid-state NMR spectroscopy [119]. In addition, even MAS NMR spectroscopy of nanoliter samples became feasible: one approach uses a conventional rotor as a carrier of a capillary containing the nanoliter-sized sample (see Fig. 9) [121–123]. Despite this extremely small sample volume, the sensitivity is even sufficient for 2D experiments [122] as well as quadrupolar tensor determination [123]. Another approach is the magic-angle coil spinning (MACS) experiment [124]. A tuned microcoil is tightly wound around the sample, which is placed in a glass capillary. The capillary and the microcoil are then inserted into a conventional MAS rotor. Wireless coupling between the tuned microcoil and the probe generates a high radio-frequency field and leads to enhanced detection sensitivity.
Spectral resolution
As already stated, solid-state NMR spectroscopy usually relies on the application of line-narrowing techniques, in particular MAS. The spectral resolution is, therefore, determined by the residual line width of the MAS-narrowed signals. Apart from trivial experimental imperfections such as a misadjustment of the magic angle, field inhomogeneities, etc., the residual line width of MAS NMR signals is determined by the following effects [125–131]:
-
1.
The incomplete suppression of the line-broadening influence of internal magnetic interactions. This is particularly true for the influence of strong homonuclear dipolar couplings (e.g., among 1H or 19F nuclei) as well as second-order quadrupolar effects for I > 1/2.
-
2.
The interfering effect of random molecular motions with the coherent averaging techniques of MAS and heteronuclear decoupling [129–131]. This effect is demonstrated in Fig. 10.
-
3.
Distributions of the isotropic chemical shift which may be caused by distributions of parameters such as bond lengths, bond angles, etc.
-
4.
Distribution of the isotropic value of the magnetic susceptibility and line-broadening due to the anisotropy of the magnetic susceptibility.
In particular the effects described in item 1 are strongly influenced by the external magnetic field B 0 as well as the sample spinning rate ν r. To evaluate the influence of B 0 and ν r on the spectral resolution, it should be remembered that the frequency difference between two neighboring signals of given chemical shift difference ∆δ increases linearly. That means the resolution increases linearly if the residual line width of the two signals remains constant at the frequency scale, i.e., if the magnetic interactions responsible for the residual line width do not depend on B 0. This is, in principle, true for the magnetic dipole–dipole interaction. Strong homonuclear magnetic dipole–dipole interactions among magnetically equivalent nuclei can only be averaged out completely by very high sample spinning rates [126–128] or in CRAMPS experiments (see earlier). The residual line width of spin systems such as 1H is dominated by the homonuclear magnetic dipole–dipole interaction and can be written as follows:
∆ν II denotes the homonuclear contribution to the static line width, i.e., the line width without MAS. The geometry-dependent factor A is typically found within the range 10–40 [126]. That means the residual line width is proportional to 1/ν r (see also Fig. 11) and the suppression of the homonuclear magnetic dipole–dipole interaction would require sample spinning far beyond 100 kHz [126, 127], which is not yet possible. This is the reason why directly detected 1H MAS NMR spectra of such systems are often poorly resolved. It is, however, important to note that the spectral resolution in homonuclear spin systems may increase even more strongly than linear in B 0. The reason for this unexpected behavior is the truncation of the Hamiltonian in homonuclear spin systems at high fields resulting in a reduction of the line width with increasing B 0 even at the frequency scale [132].
The past decade also saw considerable progress in the development of faster MAS probes [128]. Maximum sample spinning rates of approximately 70 kHz are now available. Combined with suitable homonuclear decoupling techniques [133, 134], such high sample spinning rates result in solid-state 1H NMR spectra of considerable spectral resolution.
Another possibility to reduce the homonuclear magnetic dipole–dipole interaction in strongly coupled 1H spin systems such as solid proteins is partial deuteration, a method which allows the measurement of highly resolved 1H-detected spectra of proteins in the solid state. This is of special interest: Multidimensional solid-state NMR spectra of proteins are usually not 1H-detected owing to the rather poor spectral resolution. For sensitivity reasons, however, 1H detection is highly desirable (see earlier) and, hence, commonly applied in multidimensional liquid-state NMR spectroscopy of proteins. At high levels of deuteration, extremely well resolved solid-state 1H NMR spectra of proteins can be detected [135]. The optimum exchange level of exchangeable protons in solid proteins amounts to 30–40%, as could be measured recently [136] (see also Fig. 12). HETCOR experiments on crystalline, perdeuterated proteins allow the identification of hydroxyl protons, the study of their hydrogen bonds, and determination of exchange dynamics [137].
The line width and shape of nuclei with I > 1/2 is usually determined by the electric quadrupole interaction in second-order perturbation theory [138]. This broadening is proportional to 1/B 0; therefore, increasing field strengths lead to a corresponding nonlinear improvement of spectral resolution.
Finally, it should be noted that the line width increases linearly with B 0 at the frequency scale if the residual line width reaches the limiting “natural” line width determined by the distribution of the isotropic chemical shift and/or the magnetic susceptibility of the sample. Increasing magnetic fields do then not further improve spectral resolution.
Conclusions
Triggered by continuous methodical advances, solid-state NMR spectroscopy is increasingly important for the study of biological problems. Apart from stronger magnetic fields and faster MAS devices, remarkable methodical improvements are, for example:
-
Hyperpolarization techniques (DNP, CIDNP/photo-CIDNP, SEOP)
-
Microcoils in combination with MAS (nanoliter samples!)
-
T1 shortening for sensitivity improvement.
Prominent biological applications of solid-state NMR spectroscopy are:
-
Membrane proteins/peptides
-
Protein aggregates such as amyloid fibrils
-
Photobiochemical processes (e.g., by photo-CIDNP experiments)
-
Investigations of integer cells
-
Biomineralization phenomena
Solid-state NMR spectroscopy can serve as an analytical tool to identify substances by characteristic spectroscopic parameters (chemical shifts, J-coupling constants, etc.). Furthermore, solid-state NMR spectroscopy allows investigations of the spatial structure as well as dynamics of biomolecules without the need for crystalline samples. The capability of characterizing dynamical processes is the reason why solid-state NMR spectroscopy is often useful even for crystalline samples because the provided information complements the structural information obtained by diffraction studies. In summary, it can be stated that solid-state NMR spectroscopy is an increasingly important tool in bioanalytics.
References
Wüthrich K (1986) NMR of proteins and nucleic acids. Wiley, New York
Cavanagh J, Fairbrother WJ, Palmer AG III, Skelton NJ (1996) Protein NMR spectroscopy, principles and practice. Academic, San Diego
Castellani F, van Rossum B, Diehl A, Schubert M, Rehbein K, Oschkinat H (2002) Nature 420:98
Baldus M (2007) J Biomol NMR 39:73
Andronesi OC, Becker S, Seidel K, Heise H, Young HS, Baldus M (2005) J Am Chem Soc 127:12965
Tycko R (2006) Methods Enzymol 413:103
Müller SD, Angeliss AA, Walther TH, Grage SL, Lange C, Opella SJ, Ulrich AS (2007) Biochim Biophys Acta 1768:3071
Laage S, Marchetti A, Sein J, Pierattelli R, Sass HJ, Grzesiek S, Lesage A, Pintacuda G, Emsley L (2008) J Am Chem Soc 130:17216
Manolikas T, Herrmann T, Meier BH (2008) J Am Chem Soc 130:3959
Strandberg E, Kanithasen N, Tiltak D, Bürck J, Wadhwani P, Zwernemann O, Ulrich AS (2008) Biochemistry 47:2601
Wadhwani P, Bürck J, Strandberg E, Mink C, Afonin S, Ulrich AS (2008) J Am Chem Soc 130:16515
Aisenbrey C, Bechinger B, Gröbner G (2008) J Mol Biol 375:376
Ader C, Schneider R, Hornig S, Velisetty P, Wilson EM, Lange A, Giller K, Ohmert I, Martin-Eauclaire MF, Trauner D, Becker S, Pongs O, Baldus M (2008) Nat Struct Mol Biol 15:605
Park SH, Loudet C, Marassi FM, Dufourc EJ, Opella SJ (2008) J Magn Reson 193:133
Ader C, Schneider R, Seidel K, Etzkorn M, Becker S, Baldus M (2009) J Am Chem Soc 131:170
Ader C, Schneider R, Hornig S, Velisetty P, Vardanyan V, Giller K, Ohmert I, Becker S, Pongs O, Baldus M (2009) EMBO J 28:2825
Paravastu AK, Qahwash I, Leapman RD, Meredith SC, Tycko R (2009) Proc Natl Acad Sci USA 106:7443
McDermott A (2009) Annu Rev Biophys 38:385
Sani MA, Keech O, Gardeström P, Dufourc EJ, Gröbner G (2009) FASEB J 23:2872
Resende JM, Moraes CM, Munhoz VH, Aisenbrey C, Verly RM, Bertani P, Cesar A, Pilo-Veloso D, Bechinger B (2009) Proc Natl Acad Sci USA 106:16639
Aisenbrey C, Bertani P, Bechinger B (2010) Methods Mol Biol 618:209
Wu CH, Das BB, Opella SJ (2010) J Magn Reson 202:127
Gröger C, Lutz K, Brunner E (2009) Prog Nucl Magn Reson Spectrosc 54:54
Gröger C, Sumper M, Brunner E (2008) J Struct Biol 161:55
Brunner E, Ehrlich H, Schupp P, Hedrich R, Hunoldt S, Kammer M, Machill S, Paasch S, Bazhenov VV, Kurek D, Arnold T, Brockmann S, Ruhnow M, Born R (2009) J Struct Biol 168:539
Brunner E, Richthammer P, Ehrlich H, Paasch S, Simon P, Ueberlein S, van Pée K-H (2009) Angew Chem Int Ed 48:9724
Purcell EM, Torrey HC, Pound RV (1946) Phys Rev 69:37
Bloch F, Hansen WW, Packard M (1946) Phys Rev 70:474
Pake GE (1948) J Chem Phys 16:327
Mehring M (1983) Principles of high-resolution NMR in solids, 2nd edn. Springer, Berlin
Andrew ER, Bradbury A, Eades RG (1958) Nature 182:1659
Waugh JS, Huber LM, Haeberlen U (1968) Phys Rev Lett 20:180
Haeberlen U, Waugh JS (1968) Phys Rev 175:453
Pembleton RG, Ryan LM, Gerstein BC (1977) Rev Sci Instrum 48:1286
Scheler G, Haubenreisser U, Rosenberger H (1981) J Magn Reson 44:134
Bielecki A, Kolbert AC, Levitt MH (1989) Chem Phys Lett 155:341
Vinogradov E, Madhu PK, Vega S (1999) Chem Phys Lett 314:443
Sakellariou D, Lesage A, Hodgkinson P, Emsley L (2000) Chem Phys Lett 319:253
Pines A, Waugh JS, Gibby MG (1972) J Chem Phys 56:1776
Pines A, Gibby MG, Waugh JS (1973) J Chem Phys 59:569
Metz G, Wu X, Smith SO (1994) J Magn Reson A 110:219
Hediger S, Meier BH, Kurur ND, Bodenhausen G, Ernst RR (1994) Chem Phys Lett 223:283
Hediger S, Meier BH, Ernst RR (1995) Chem Phys Lett 240:449
Bennett AE, Rienstra CM, Auger M, Lakshmi KV, Griffin RG (1995) J Chem Phys 103:6951
Fung BM, Khitrin AK, Ermolaev K (2000) J Magn Reson 142:97
Iuga A, Ader C, Gröger C, Brunner E (2007) Annu Rep NMR Spectrosc 60:145
Gullion T, Schaefer J (1989) J Magn Reson 81:196
Levitt MMH, Raleigh DP, Creuzet F, Griffin RG (1990) J Chem Phys 92:6347
Ernst RR, Bodenhausen G, Wokaun A (1987) Principles of nuclear magnetic resonance in one and two dimensions. Clarendon Press, Oxford
Bax A, Freeman R, Kempsell SP (1980) J Am Chem Soc 102:4849
Levitt MH, Ernst RR (1983) Mol Phys 50:1109
Lesage A, Bardet M, Emsley L (1999) J Am Chem Soc 121:10987
Baldus M, Meier BH (1996) J Magn Reson A 121:65
Fyfe CA, Wong-Moon KC, Huang Y, Grondey H (1995) J Am Chem Soc 117:10397
Elena B, Lesage A, Steuernagel S, Böckmann A, Emsley L (2005) J Am Chem Soc 127:17296
Mao K, Pruski M (2009) J Magn Reson 201:165
Scheidt HA, Schibur S, Magalhães A, de Azevedo ER, Bonagamba TJ, Pascui O, Schulz R, Reichert D, Huster D (2010) Biopolymers 93:520
Krushelnitsky A, de Azevedo E, Linser R, Reif B, Saalwächter K, Reichert D (2009) J Am Chem Soc 131:12097
Li W, McDermott A (2009) J Biomol NMR 45:227
Abragam A (1961) Principles of nuclear magnetism. Oxford University Press, Oxford
Hoult DI, Richards RE (1976) J Magn Reson 24:71
Kozlov MB, Haase J, Baumann C, Webb AG (2005) Solid State Nucl Magn Reson 28:64
Spiess HW (2008) Angew Chem Int Ed 47:639
Opella SJ (2009) Nat Meth 6:197
Abragam A, Goldman M (1978) Rep Prog Phys 41:395
Matsuki Y, Maly T, Ouari O, Karoui H, Le Moigne F, Rizzato E, Lyubenova S, Herzfeld J, Prisner T, Tordo P, Griffin RG (2009) Angew Chem Int Ed 48:4996
Barnes AB, De Paëpe G, van der Wel PCA, Hu K-N, Joo C-G, Bajaj VS, Mak-Jurkauskas ML, Sirigiri JR, Herzfeld J, Temkin RJ, Griffin RG (2008) Appl Magn Reson 34:237
Maly T, Debelouchina GT, Bajaj VS, Hu K-N, Joo C-G, Mak-Jurkauskas ML, Sirigiri JR, van der Wel PCA, Herzfeld J, Temkin RJ, Griffin RG (2008) J Chem Phys 128:052211
Rosay M, Tometich L, Pawsey S, Bader R, Schauwecker R, Blank M, Borchard PM, Cauffman SR, Felch KL, Weber RT, Temkin RJ, Griffin RG, Maas WE (2010) Phys Chem Chem Phys (in press). doi:10.1039/c003685b
Van der Wel PCA, Hu K-N, Lewandowski J, Griffin RG (2006) J Am Chem Soc 128:10840
Barnes AB, Andreas LB, Huber M, Ramachandran R, van der Wel PCA, Veshtort M, Griffin RG, Mehta MA (2009) J Magn Reson 200:95
Bajaj VS, Mak-Jurkauskas ML, Belenky M, Herzfeld J, Griffin RG (2010) J Magn Reson 202:9
Bargon J, Fischer H (1967) Z Naturforsch 22a:1556
Ward HR, Lawler RG (1967) J Am Chem Soc 89:5518
Lawler RG (1967) J Am Chem Soc 89:5519
Bargon J (2006) Phys Chem Chem Phys 5:970
Cocivera M (1968) J Am Chem Soc 90:3261
Kaptein R, den Hollander JA, Antheunis D, Oosterhoff LJ (1970) J Chem Soc D 24:1687
Zysmilich MG, McDermott A (1994) J Am Chem Soc 116:8362
Schulten EAM, Matysik J, Alia, Kiihne S, Raap J, Lugtenburg J, Gast P, Hoff AJ, de Groot JM (2002) Biochemistry 41:8708
Roy E, Rohmer T, Gast P, Jeschke G, Alia A, Matysik J (2008) Biochemistry 47:4629
Diller A, Roy E, Gast P, van Gorkom HJ, de Groot HJM, Glaubitz C, Jeschke G, Matysik J, Alia A (2007) Proc Natl Acad Sci USA 104:12767
Daviso E, Prakash S, Alia A, Gast P, Neugebauer J, Jeschke G, Matysik J (2009) Proc Natl Acad Sci USA 106:22281
Walker T, Happer W (1997) Rev Mod Phys 69:629
Appelt S, Baranga B-A, Erickson CJ, Romalis MV, Young AR (1998) Phys Rev A 58:1412
Fink A, Baumer D, Brunner E (2005) Phys Rev A 72:053411
Ito T, Fraissard J (1982) J Chem Phys 76:5225
Raftery D, Long H, Meersmann T, Grandinetti PJ, Reven L, Pines A (1991) Phys Rev Lett 66:584
Bowers CR, Storhaug V, Webster CE, Bharatam J, Cottone A 3rd, Gianna R, Betsey K, Gaffney BJ (1999) J Am Chem Soc 121:9370
Locci E, Dehouck Y, Casu M, Saba G, Lai A, Luhmer M, Reisse J, Bartik K (2001) J Magn Reson 150:167
Rubin SM, Spence M, Goodson BM, Wemmer DE, Pines A (2000) Proc Natl Acad Sci USA 97:9472
Rubin SM, Lee S-Y, Ruiz EJ, Pines A, Wemmer DE (2002) J Mol Biol 322:425
Gröger C, Möglich A, Pons M, Koch B, Hengstenberg W, Kalbitzer HR, Brunner E (2003) J Am Chem Soc 125:8726
Dubois L, de Silva P, Landon C, Huber JG, Ponchet M, Vovelle F, Berthault P, Desvaux H (2004) J Am Chem Soc 126:15738
Anedda R, Era B, Casu M, Fais A, Ceccarelli M, Corda M, Ruggerone P (2008) J Phys Chem B 112:15856
Brunner E (1999) Conc Magn Reson 11:313
Raftery D (2006) Annu Rep NMR Spectrosc 57:205
Albert MS, Cates GD, Driehuys B, Happer W, Saam B, Springer CS Jr, Wishnia A (1994) Nature 370:199
Matsuoka S, Patz S, Albert MS, Sun Y, Rizi RR, Gefter WB, Hatabu H (2009) J Thorac Imaging 24:181
Brunner E, Seydoux R, Haake M, Pines A, Reimer JA (1998) J Magn Reson 130:145
Navon G, Song Y-Q, Rõõm T, Appelt S, Taylor RE, Pines A (1996) Science 271:1848
Desvaux H, Marion DJ, Huber G, Dubois L, Berthault P (2006) Eur Phys J Appl Phys 36:25
Schröder L, Lowery TJ, Hilty C, Wemmer DE, Pines A (2006) Science 314:446
Berthault P, Bogaert-Buchmann A, Desvaux H, Huber G, Boulard Y (2008) J Am Chem Soc 130:16456
Schlundt A, Kilian W, Beyermann M, Sticht J, Günther S, Höppner S, Falk K, Roetzschke O, Mitschang L, Freund C (2009) Angew Chem 121:4206
Bowers CR, Weitekamp DR (1986) Phys Rev Lett 57:2645
Eisenschmid TC, Kirss RU, Deutsch PP, Hommeltoft SI, Eisenberg R, Bargon J, Lawler RG, Balch AL (1987) J Am Chem Soc 109:8089
Pravica M, Weitekamp DP (1988) Chem Phys Lett 145:255
Korchak SE, Ivanov KL, Yurkovskaya AV, Vieth H-M (2009) Phys Chem Chem Phys 11:11146
Haake M, Natterer J, Bargon J (1996) J Am Chem Soc 118:8688
Kuhn LT, Bommerich U, Bargon J (2006) J Phys Chem A 110:3521
Carson PJ, Bowers CR, Weitekamp DP (2001) J Am Chem Soc 123:11821
Adams RW, Aguilar JA, Atkinson KD, Cowley MJ, Elliott PIP, Duckett SB, Green GGR, Khazal IG, López-Serrano J, Williamson DC (2009) Science 323:1708
Mishkovsky M, Frydman L (2008) Chemphyschem 9:2340
Wickramasinghe NP, Kotecha M, Samoson A, Past J, Ishii Y (2007) J Magn Reson 184:350
Linser R, Chevelkov V, Diehl A, Reif B (2007) J Magn Reson 189:209
Wickramasinghe NP, Parthasarathy S, Jones CR, Bhardwaj C, Long F, Kotecha M, Mehboob S, Fung LW-M, Past J, Samoson A, Ishii Y (2009) Nat Meth 6:215
Lopez JJ, Kaiser C, Asami S, Glaubitz C (2009) J Am Chem Soc 131:15970
Yamauchi K, Janssen JWG, Kentgens APM (2004) J Magn Reson 167:87
Kentgens APM, Bart J, van Bentum PJM, Brinkmann A, van Eck ERH, Gardeniers JGE, Janssen JWG, Knijn P, Vasa S, Verkuijlen MHW (2008) J Chem Phys 128:052202
Janssen H, Brinkmann A, van Eck ERH, van Bentum JM, Kentgens APM (2006) J Am Chem Soc 126:8722
Brinkmann A, Vasa SK, Janssen H, Kentgens APM (2010) Chem Phys Lett 485:275
Vasa SK, van Eck ERH, Janssen JWG, Kentgens APM (2010) Phys Chem Chem Phys 12:4813
Sakellariou D, Le Goff G, Jacquinot J-F (2007) Nature 447:694
Maricq MM, Waugh JS (1979) J Chem Phys 70:3300
Brunner E, Freude D, Gerstein BC, Pfeifer H (1990) J Magn Reson 90:90
Brunner E (1990) J Chem Soc Faraday Trans 86:3957
Samoson A, Tuherm T, Past J, Reinhold A, Anupõld T, Heinmaa I (2004) Top Curr Chem 246:15
Suwelack D, Rothwell WP, Waugh JS (1980) J Chem Phys 73:2559
Rothwell WP, Waugh JS (1981) J Chem Phys 74:2721
Fenzke D, Gerstein B, Pfeifer H (1992) J Magn Reson 98:469
van Rossum B-J, Boender GJ, de Groot HJM (1996) J Magn Reson A 120:274
Leskes M, Steuernagel S, Schneider D, Madhu PK, Vega S (2008) Chem Phys Lett 466:95
Lafon O, Wang Q, Hu B, Trébosc J, Deng F, Amoureux J-P (2009) J Chem Phys 130:014504
Chevelkov V, Rehbein K, Diehl A, Reif B (2006) Angew Chem Int Ed 45:3878
Akbey Ü, Lange S, Franks WT, Linser R, Rehbein K, Diehl A, van Rossum B-J, Reif B, Oschkinat H (2010) J Biomol NMR 46:67
Agarwal V, Linser R, Fink U, Faelber K, Reif B (2010) J Am Chem Soc 132:3187
Ashbrook SE (2009) Phys Chem Chem Phys 11:6892
Acknowledgements
The authors wish to thank Annett Bachmann and Renate Schulze for carefully proofreading the manuscript. Financial support from the Deutsche Forschungsgemeinschaft (BR 1278/ 12) is gratefully acknowledged.
Author information
Authors and Affiliations
Corresponding author
Additional information
Published in the special issue on Focus on Bioanalysis with Guest Editors Antje J. Baeumner, Günter Gauglitz, and Frieder W. Scheller.
Rights and permissions
About this article
Cite this article
Paasch, S., Brunner, E. Trends in solid-state NMR spectroscopy and their relevance for bioanalytics. Anal Bioanal Chem 398, 2351–2362 (2010). https://doi.org/10.1007/s00216-010-4037-5
Received:
Revised:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1007/s00216-010-4037-5