Isotope-Aided Methods for Biological NMR Spectroscopy: Past, Present, and Future
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This chapter starts by providing a historical background of our research endeavors over the past half-century to develop various isotope-aided methods in biological NMR spectroscopy, since innovations bloom only on the rich ground cultivated by previous investigators. We then focused on the stereo-array isotope-labeling (SAIL) method, one of our recent accomplishments, which culminates the isotope-aided NMR technologies for structural studies of proteins from various aspects: accurate structural determinations of large proteins, elaboration for automated structural determination, highly efficient and versatile residue-selective methyl labeling with newly developed auxotrophic E. coli strains, large-amplitude slow-breathing motion (LASBM) as revealed by the aromatic ring flipping of the residues in ligand-binding interfaces, and applications of the deuterium-induced 13C-NMR isotope shift to investigate the hydrogen exchange phenomena of side-chain polar groups. Meanwhile, the expected role of NMR spectroscopy has been rapidly shifting from structure determinations to dynamics studies of biologically interesting targets, such as membrane proteins and larger protein complexes. The dynamic aspects of protein–protein and protein–ligand interactions are closely related to their biological functions and can be efficiently studied by using proteins residue selectively labeled with amino acids bearing optimized labeling patterns, prepared by cellular expression. We are absolutely confident that biological NMR spectroscopy will continually develop with further innovations of isotope-labeling technologies in the coming era, featuring ultrahigh field spectrometers beyond 1 GHz.
KeywordsIsotope-aided NMR method 13C, 15N double labeling SAIL method Aromatic ring flipping Stereo-specific methyl labeling Hydrogen exchange rates of polar groups Deuterium-induced 13C isotope shift
2.1 Historical Background of Our Research to Develop Isotope-Aided NMR Methods
Over the past half-century, we have exploited isotope-aided NMR methods to investigate biological molecules, such as amino acids, peptides, proteins, and nucleic acids. Many of our results were initially presented at the annual meetings of the NMR Society of Japan, and some of our novel techniques are still being used worldwide. One of the prominent features of our approach is utilizing site- and stereo-specifically isotope-labeled amino acids and nucleosides, which can be efficiently prepared by combining microbial fermentations, enzymatic reactions, and chiral organic syntheses—all areas in which Japan has world-leading technological expertise. In this chapter, after a brief recollection of the early days in the development of isotope-aided biological NMR methods, we describe some of the past and current advances achieved mostly in our laboratory. However, because of the space limitation, the methods for studying nucleic acids are not included.
At the first annual symposium of the NMR Society of Japan, which was held in Tokyo, 1961, two papers on NMR studies of amino acids were presented. Namely, Fujiwara et al. reported the 56.4 MHz 1H-NMR spectra of aqueous solutions of various amino acids and Takeuchi et al. reported the 40 MHz 1H-NMR spectra of threonine and allo-threonine. To the best of our knowledge, these were the first biological NMR applications ever reported in Japan. At that time, there was no systematic NMR research on proteins, except for a short communication on ribonuclease A . Unfortunately, the reported 40 MHz spectrum showed only four broad overlapped signals and did not provide detailed structural information. It was obvious that some groundbreaking methodologies were absolutely necessary to investigate proteins by NMR. Jardetzky proposed an excellent idea at the “International Symposium on Nuclear Magnetic Resonance,” which was held in Tokyo, 1965, aiming to settle the problem . In this very exciting international meeting, gathering many of the eminent NMR pioneers, he explained a perspective of biosynthetic selective deuteration to simplify the 1H-NMR spectra of proteins for obtaining structural information related to their biological functions. Surprisingly, it was only three years later when his colleagues published the first 100 MHz 1H-NMR spectra of selectively deuterated nucleases, clearly showing that the proposed strategy actually works well . This enlightening work strongly motivated us to further develop isotope-labeling technologies for NMR studies of biological systems ever since .
2.1.1 Stereo-Specific Deuteration of Prochiral Methylene Protons—Conformational Analysis of Amino Acids and Peptides
2.1.2 Selective 13C, 15N Double-Labeling Method for the Sequential Assignment of Backbone Amide NMR Signals in Large Proteins
During the 1970s, there was a multidisciplinary collaborative research group, known as the “Research Consortium on Streptomyces Subtilisin Inhibitor (SSI),” which focused on SSI as a shared target, aiming to promote biophysical and biochemical protein research activities in Japan. SSI, which was isolated from the culture broth of Streptomyces albogriseolus, is a 23-kDa dimeric protein composed of two identical subunits, and it strongly inhibits serine proteinases, especially subtilisin family proteinases. One of the controversial issues for proteinase–inhibitor interactions in the late 1970s was the state of the active site peptide bond, i.e., the “scissile bond,” in the inhibitors complexed with proteinases. X-ray crystallographic analyses of various proteinase–inhibitor complexes initially concluded the existence of the “tetrahedral intermediate,” which was thought to be formed by a covalent bond between the active site Ser Oγ of a proteinase and the carbonyl of a scissile bond. It was assumed that the tetrahedral intermediate was trapped due to the overwhelming stabilization by the “oxy-anion hole” of serine proteinases, which serves as a molecular device to enormously accelerate the enzymatic hydrolysis of substrate peptides. However, this remarkable model, which was cited in most of the biochemistry textbooks at that time to highlight the beauty of the enzymatic functions of proteinases, was becoming dubious. Namely, as higher-resolution X-ray structures became available, they revealed that the atomic distance between the Ser Oγ and the carbonyl carbon atom of the scissile bond was apparently a little too long to form the covalent bond. In addition to this serious concern, others still remained about the state of the proteinase–inhibitor complex, which could not be solved by crystallographic studies, such as the following: Does it exist in solution as a single intermediate or as an equilibrium mixture of multiple intermediates? Is the scissile peptide bond in the complexes planar, as usually found for peptide bonds, or distorted by the effect of the nearby Ser Oγ? In principle, all of these questions could be investigated by solution NMR spectroscopy. However, it was difficult to think that such work was feasible since the molecular weight of the SSI–subtilisin complex, 78 kDa, was too large. We overcame this problem by a unique isotope-labeling strategy, as described below .
The idea of the sequential assignment for the backbone carbonyl carbons by the selective 13C, 15N double-labeling method was extended to assign the peptide 15N and side-chain 13C signals, through the 13C–15N and 13C–13C connectivities [10, 11]. The method can be regarded as the historic prototype of triple-resonance sequential assignment methods, using uniformly 13C, 15N double-labeled proteins. We were actually one of the first groups to suggest the idea of establishing sequential assignment methods by extending the 13C, 15N double-labeling method, which was presented at the XI ICMRBS in Goa, India, in 1984 .
2.1.3 Revisiting the Stereo-Specific Isotope-Labeling Approach for Studying Proteins: A Long March to the SAIL Method
In 1997, with financial support from the newly launched grant “Core Research and Evolutional Science and Technology,” also known as CREST, aiming to promote basic sciences in Japan, we started a 5-year project to develop a breakthrough isotope-aided NMR technology for studying large proteins. At that time, NMR technologies using uniformly 13C, 15N double-labeled proteins were firmly established, but they could only be applied to determine the three-dimensional structures of small proteins. However, a variety of key technologies, which were not available back in the 1970s, facilitated further innovations of isotope-aided NMR methods. For example, multinuclear multidimensional NMR spectroscopy, chiral organic syntheses, and protein expression using recombinant DNA techniques were all quite mature techniques by then. Therefore, we had a unique opportunity to revisit the old idea to explore cutting-edge methods for studying larger proteins. In order to encompass the advents of various multidimensional NMR methods, we exploited novel synthetic routes for the regio- and stereo-specifically D, 13C, 15N triple-labeled amino acids. With the help of state-of-the-art chiral organic synthetic methods, together with enzymatic reactions and microbial fermentations, we actually successfully synthesized all of the protein component amino acids with a variety of labeling patterns [13, 14, 15]. Fortunately, with further support from a second CREST grant for another five years, we completed the development of the “stereo-array isotope-labeling (SAIL) method” by using those labeled amino acids. Although it took more than 30 years after our early work on the stereo-specific deuteration of amino acids and peptides, the SAIL method has been proven to be extremely useful for studying the structures as well as the dynamics of larger proteins, for which previous NMR methods were difficult to apply [16, 17, 18, 19].
2.2 The SAIL Method: An Optimized Isotope-Labeling Strategy for the Structural Study of Proteins by NMR Spectroscopy
NMR spectra of larger proteins are typically characterized by numerous overlapped signals, which are severely broadened by dipolar interactions between nearby protons. Therefore, it was difficult to obtain sufficient amounts of NMR information for proteins larger than 20–25 kDa, even with sophisticated multidimensional methods. For example, for a long time it was thought to be virtually impossible to analyze the prochiral methylene proton signals, especially for large proteins, even though the information is absolutely required for accurately determining the side-chain conformations, as described above for amino acids [5, 6]. Conceptually, however, we may not necessarily need all of the NMR data for the determination of protein structures, since many of the amino acid side chains contain somewhat redundant information. For example, if we could stereo-specifically observe either one of the prochiral groups, i.e., methyls and methylenes, we could compensate for the missing information about the geminal counterparts. The SAIL method creates this type of situation for all of the amino acid residues in proteins, by trimming away the redundant information by the optimized isotope-labeling patterns, as described below [20, 21].
2.2.1 Cell-Free Expression and NMR Spectra of SAIL Proteins
As described above, the SAIL method facilitates structural analyses of larger proteins, through the rational isotope-labeling design for the component amino acid residues. Therefore, it is necessary to incorporate SAIL amino acids into a target protein while preserving their original labeling patterns. In this respect, conventional cellular protein expression using recombinant DNA may not be a good choice, since metabolic scrambling reactions and isotope dilution with unlabeled amino acids are not completely avoidable for some amino acid residues. All these problems are largely circumvented by using in vitro expression systems. Fortunately for us, the in vitro protein expression using the E. coli cell-free extract became available at around the time we nearly completed the synthesis of the SAIL amino acids. By using the cell-free extract prepared from the E. coli cells, we successfully prepared a sufficient amount of a SAIL protein for an NMR study. SAIL proteins, which are composed exclusively of SAIL amino acids, are typically obtained with ~10 wt% yields calculated from the amount of the amino acid mixture and show virtually no isotopic dilution or metabolic scrambling [23, 24, 25].
2.2.2 Structural Determination of SAIL Proteins
The marked improvement in the overall quality of the NMR spectra obtained for SAIL proteins further encouraged us to use the FLYA program for automated backbone and side-chain resonance assignments. The chemical shift data obtained by FLYA can then be used as the input data for the NOESY spectral analysis and the structure calculation by CYANA. This two-step automated structure determination using the FLYA-CYANA program works well for small SAIL proteins, without additional human participation [29, 30]. We also tried to develop a fully automated structure determination method exclusively based on NOESY data, obviating the need to measure any other spectra than those necessary for the resonance assignment. This ambitious automated approach, which would be useful for determining a large number of structures as efficiently as possible, was actually utilized for two small SAIL proteins and yielded well-defined structures that coincide closely with those determined by the conventional method .
2.3 Recent Trends in the Isotope-Aided NMR Methods for Studying Proteins
We described above our early studies on isotope-aided NMR techniques and then introduced a recent achievement, the SAIL method. However, the prospective roles of NMR in structural biology are rapidly changing, especially because other methods, such as X-ray crystallography and cryo-electron microscopy, have been overwhelmingly employed for the structural determinations of biologically important proteins, such as membrane proteins or extraordinarily large protein complexes. Obviously, NMR cannot be a competitive structural determination tool for those targets. Instead, a variety of alternative applications are envisaged for NMR spectroscopy, to bridge the gap between protein structures and their biological functions. In principle, NMR could afford unique information for this purpose, even if the proteins are too large for structure determination by NMR. Actually, in many cases, one could start with the three-dimensional structures previously determined by the other methods and focus on the structures and dynamics of the selected regions of interest, which could be precisely manifested by NMR. For that purpose, it is necessary to develop a method to observe and assign the NMR signals for any regions of the selected amino acid residues in such proteins. Recently, there have been major advances in NMR signal observations for larger proteins. Wuethrich et al. exploited transverse relaxation-optimized spectroscopy (TROSY) for observing the backbone amide 1H15N signals in deuterated proteins and the aromatic ring 1H13C signals for uniformly 13C-labeled proteins [32, 33]. Kay et al. developed a method to observe the Ile, Leu, and Val (ILV) 13CH3 signals, utilizing them as NMR probes for studying protein structures and dynamics . The 1H15N TROSY and methyl observation methods can be applied for proteins as large as 1 MDa and are routinely used for studying larger proteins in solution .
Since the backbone amides and the side-chain methyl groups (13CH3) cover considerable portions of larger proteins, their NMR signals provide valuable structural information. However, it may not be sufficient for analyzing the precise side-chain conformations and dynamics for the selected residues in order to understand the molecular basis of biological functions, which we expect to obtain by solution NMR. This is where sophisticated isotope-aided methods such as SAIL come in. We have exploited methods to observe NMR signals for any parts of aliphatic and aromatic side chains in a protein, by further optimizing the isotope-labeling patterns of the original SAIL design concepts. In the following, we describe some of our recent work along this line, in order to provide an outline of our current research endeavors.
2.3.1 Residue- and Stereo-Specific Labeling Method: The Case for Leu and Val Methyl Labeling of Larger Proteins
Larger proteins have numerous methyl groups in their Ala, Thr, Met, Ile, Leu, and Val (ATMILV) residues, which are widely distributed on their surface and interior regions. Therefore, the methyl signals are valuable probes for studying the structures and dynamics of proteins and protein complexes, if the individual methyl signals could be observed. However, this is not trivial especially for larger proteins, since they have so many ATMILV residues. For example, the 82-kDa protein malate synthase G (MSG) has 289 ATMILV residues that comprise approximately 40% of its 723 residues. Among them, the 160 ILV residues are especially useful as NMR probes, since the total of 317 ILV methyls accounts for as many as ~70% of the 446 methyl signals. Therefore, extensive efforts have been exerted to develop robust methods to observe and assign ILV methyl signals in larger proteins. Most of them employ region-specifically 13C, D-labeled precursors, such as [4-13C;3,3-D2]-α-ketobutyrate and [3-13CH3;3,4,4,4-D4]-α-ketoisovalerate (α-KIV), for preparing fully deuterated proteins except for the Ile (δ1), Val (γ1/γ2), and Leu (δ1/δ2) methyls, which are to be labeled with 13CH3 [36, 37]. However, since the racemic α-KIV precursor labels both of the prochiral methyls in Leu and Val residues, the observable number of methyl signals cannot be decreased, and the labeling rates are 50% or less. Therefore, the signal congestion for Leu/Val methyls could not be improved and the methyl–methyl NOEs are significantly reduced. In order to compensate for the drawbacks of this Leu/Val precursor, a few new precursors for the stereo-specific methyl labeling of Leu/Val residues have been developed . However, it is difficult to use labeled precursors to prepare any desired combinations for either one of the prochiral methyls in Val and Leu, since Val is converted to Leu biosynthetically, as shown in the metabolic map.
2.3.2 Large-Amplitude Dynamics of Proteins as Probed by Aromatic Ring-Flipping Motions—The Case for the Interface Between FKBP and Drug Complexes
Nowadays, it is generally accepted that folded proteins occasionally undergo large-amplitude slow-breathing motions (LASBMs) under physiological conditions. Since the frequency of LASBMs is within the millisecond to second region, such motions have attracted the interests of biophysicists in the context of biological functions and protein dynamics. The LASBM was initially implicated by the intriguing observations that the δ- and ε-protons for the Phe and Tyr aromatic rings in proteins showed time-averaged NMR signals. It was quite surprising, especially for most of the crystallographers back in the 1970s, that such bulky aromatic rings flip about the Cβ–Cγ axis so frequently, since they are often deeply embedded in the hydrophobic core, which was thought to be the most solid part of a protein [41, 42]. Ironically, until recently there have only been few cases in which the aromatic rings show discrete signals for the δ- and ε-nuclei of Phe and Tyr, due to slow ring-flipping rates . Theoretically, it might be possible that the 1H and 13C nuclei at the δ- and ε-positions incidentally have identical, or nearly identical, chemical shifts and thus appear as a time-averaged single peak regardless of the flipping rates, although it is quite unlikely that such situations happen very often. Taking advantage of the simplified spin systems of the SAIL Phe and Tyr, we revisited the ring-flipping phenomena and found that there are actually many more cases showing flipping rate-dependent aromatic ring signals. Apparently, the aromatic rings in conventional protein samples have such complicated spin networks that such cases are rarely identified. Therefore, proteins selectively labeled by δ-, ε-SAIL Phe and Tyr would provide unprecedented opportunities to investigate LASBMs through the widely distributed aromatic rings in the hydrophobic interior and on the ligand-binding surface. In the following, we illustrate the application of the aromatic ring-flipping phenomena for characterizing the LASBMs within the binding interface in FKBP12–ligand complexes .
2.3.3 Deuterium-Induced Isotope Shifts for Measuring Hydrogen Exchange Rates of Polar Side-Chain Groups in Proteins: Facile Screening of the Polar Groups Involved in Hydrogen Bond Networks
The hydrogen exchange phenomena of the backbone amides in aqueous solutions are among the most intensively studied protein dynamics by NMR spectroscopy. The exchange rates are usually estimated by a time course of the amide proton signal intensity changes for a protein freshly dissolved in D2O. The information has made crucial contributions toward understanding the backbone dynamics and the folding–unfolding processes of proteins in solution. In contrast, the hydrogen exchange rates for the polar side-chain groups, such as hydroxyl (OH) or sulfhydryl (SH), have not been studied extensively, because they are usually too rapid to be measured by the method used for the backbone amides. We have exploited an alternative approach for the facile screening of the slowly exchanging polar side-chain groups and the estimation of their hydrogen exchange rates with the surrounding water. We adapted our previous method for detecting slowly exchanging backbone amide hydrogens by the steady-state line shapes of the amide carbonyl 13C signals, in a protein dissolved in a 1:1 mixture of H2O and D2O [7, 8, 45]. In such an environment, the line shape of the amide carbonyl of the ith residue depends on the isotope shift values induced by deuteration for both of the (i + 1)th and ith amides and also on their hydrogen–deuterium exchange rates .
Since the 13C chemical shift differences for the carbons directly bonded to side-chain OH or SH groups measured in H2O and D2O are usually a little greater than 0.1 ppm, we could use these relatively large isotope shifts for the facile screening of the slowly exchanging polar groups. To do so, we prepared proteins selectively labeled by ζ-SAIL Tyr, [3-13C; 3,3′-D2]-Ser, [3-13C; 3-D]-Thr, or [3-13C; 3,3′-D2]-Cys. These labeled proteins gave extremely sharp 1D 13C-NMR signals for the Cζ or Cβ under deuterium decoupling and thus were quite useful for estimating the isotope shifts on the Cζ or Cβ and then the hydrogen exchange rates for the slowly exchanging polar groups by the EXSY experiment. We found that quite a few Tyr, Ser, Thr, and Cys residues in various proteins, dissolved in a 1:1 H2O–D2O mixture, actually exhibit slow hydrogen–deuterium exchanging rates for their side-chain hydroxyl or sulfhydryl groups. Interestingly, all of the polar groups identified by this method as having very slow hydrogen exchange rates form hydrogen bonds and give 1H-NMR signals in H2O [46, 47, 48, 49]. Therefore, this approach is useful for screening slowly exchanging polar functional groups that are likely to play important structural roles in proteins. A typical example of a search for the Tyr residues in a protein, which might have slowly exchanging hydroxyl groups, is illustrated as follows.
2.4 Future Perspectives of the Isotope-Aided NMR Method
The SAIL method has become well known worldwide as a state-of-the-art isotope-aided NMR technology. However, it is clear that further efforts are required to make it a standard practice among the international biological NMR communities. The substantially high cost of the SAIL amino acids is certainly one of the obstacles, but they will be more affordable if the SAIL method becomes routinely used. Cell-free protein expression, which is necessary to prepare proteins exclusively composed of SAIL amino acids, seems to be another barrier for most NMR laboratories with no such experience. However, the E. coli cell-free kit for preparing isotope-labeled protein samples for NMR is now available commercially at a moderate cost. Therefore, there are no major hurdles to trying out the SAIL method for structure studies of proteins. The SAIL method could be extended for solving precise structures of proteins as large as 100 kDa or even more, by further optimization of the relaxation properties of SAIL amino acids (Miyanoiri et al., unpublished). The applications of the SAIL method to solid-state NMR spectroscopy are also interesting, but they have just started .
Meanwhile, the expected role of NMR spectroscopy in structural biology seems to be rapidly shifting from structure determinations to dynamics studies of biologically interesting targets, such as membrane proteins and larger protein complexes. The dynamic aspects of protein–protein and protein–ligand interactions are closely related to their biological functions and can be efficiently studied by using proteins residue selectively labeled with amino acids bearing optimized labeling patterns, prepared with conventional cellular expression systems. It is therefore quite important for the NMR community to explore the stable isotope-labeling technology to its full potentials. We are absolutely confident that biological NMR spectroscopy will be continually developing with further innovations of isotope-labeling methods in the coming era, featuring ultrahigh field spectrometers beyond 1 GHz.
None of our work cited in this review could have been accomplished without the seminal contributions of numerous students and collaborators over the last half-century. We sincerely regret that we could not mention most of their important accomplishments, simply because of the space limitation. Needless to mention, our research activities were not possible without the generous, persistent financial support from various government organizations and funding agencies, although we cannot mention them individually.
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