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1 Introduction to the NMR of Proteins David G. Reid, Lesley K. MacLachlan, Andrew J. Edwards, Julia A. Hubbard, and Patricia J. Sweeney 1. Introduction There are numerous modern, and older but nonetheless valuable, textbooks that should be consulted for an introduction to the basic principles (14) and practices of nuclear magnetic resonance (NMR) in chemistry (7-19) (including volumes devoted primarily to exchanging systems [20,21], nuclei less familiar to the organic chemist [22,23], and solids [24,25]), biochemistry (2634), including parts of recent volumes in this series (35,36), and drug discovery (37). 1.1. Basic NMR Parameters The familiar NMR parameters by which organic compounds can be charac- terized are also relevant to the study of proteins. These are formally defined and well described m most of the texts already cited (such as in the first three chapters of ref. 35) and will only be mentioned briefly m this introduction. The chemical shift of a specific nucleus depends on its covalent chemistry and (usually to a lesser extent) its nonbonded environment. In particular, mvolvement m hydrogen bonding, and proximity to aromatic and carbonyl functions, cause the biggest deviations of chemical shifts from the values they would display in an unstructured, i.e., random coil, peptide. The random coil proton chemical shifts for the twenty common amino acids are shown in Table 1. Effects owing to the nonbonded environment can be a function of protein secondary and tertiary structure, and can provide useful information about the folded state. Internuclear throughbond (scalar or J) couplings can be analyzed in terms of torsion angles, and provide information about peptide backbone and side chain conformations. Relaxation times (or their reciprocal relaxation rates) are functions of molecular mobility. The spin-lattice, or longitudinal, From Methods m Molecular Biology, Vo/ 60 Protem NMR Techmques Edlted by D G Field Humana Press Inc , Totowa, NJ 2 Reid et al. Table 1 lH Chemical Shifts of Protons on Nonterminal Amino Acid Residues of Random Coil Peptides and Protein@ Residue Abbrevlatlon Ha HP Hy H6 HE HI; Ala A 4 35 1.40 A% R 4.40 1.80 1.72 3.31 1.92 Asn N 4.76 2.76 2.83 Asp D 4.77 2.75 2.84 CYS C 4 69 2 96 3 28 Gln Q 4.37 201 2 38 2.13 Glu E 4.30 1.97 2.28 2.09 231 GUY G 3.97 His H 4 63 3 20 7 14 8.12 3.26 (H4) W2) Ile I 4 22 1.89 1 19 0 89 1.48 0.94 Leu L 4.39 1.65 1.65 0.94 0.90 LYS K 4.36 1.75 147 1 71 3 02 1.87 Met M 4.51 2.00 2.63 2.13 2.16 Phe F 4.66 2 99 7 34 7.34 7 34 3.22 Pro P 471 1.98 2 30 3.65 2.30 Ser S 4.50 3.89 Thr T 4.35 4.22 1.23 Trp W 4.70 3.20 7.24 7.17 7.50 3.22 U-W (H5) (H7) 7.65 7.24 (H4) 036) Tyr Y 4.60 2 92 7.15 6.86 3 13 WV) (H3,5) Va V 4.18 2.13 0.94 0.97 aAdapted from ref. 40 Chemical shifts are cited m parts per milhon (ppm) relative to internal sodium 3-tnmethylalyl proplonate (TSP), pD = 7.0. Followmg the usual convention a downfield shift 1s posltlve. Introduction to the NMR of Proteins 3 relaxation time, T, (or rate RJ is a measure of the efficiency with which excited nuclear spins return to their ground states by exchanging energy with their surroundmgs. The spin-spm, or transverse, relaxation time, Tz (or rate, Rz) measures the efficiency with which spms exchange energy with each other. The more efficient the exchange the smaller, or shorter, the relaxation time (or the larger the relaxation rate). There are several mechamsms by which this exchange of spin energy can take place. In diamagnetic proteins the dipole- dipole interaction between spin l/2 nuclei is usually of predominant importance, but chemical shift anisotropy (CSA), nuclear quadrupolar (for nuclei with spin > l/2), and paramagnetic, effects can all be important under certain circumstances described m several subsequent chapters. An important phenomenon related to nuclear relaxation is the nuclear Overhauser effect (nOe or NOE) (38,39), formally defined as the “change in intensity of the signal of a given nucleus when the resonance of another nucleus, which contributes to its relaxation by a dipole-dipole mechanism, is saturated.” The NOE observed between two nuclei depends prmcipally on the distance between them, as well as other parameters, and hence on the folded protein conformation, if the nuclei are in the same molecule or on the structure of a complex if they are in different molecules. NMR resonance linewidths m solution are, generally speaking, inversely proportional to the T, relaxation time, which decreases with increasing molecular size and tumbling time, a measure of which is the molecular correlation time, z,. Unfortunately this usually means that as the molecular weight increases, so do the resonance linewidths, exacerbating problems of sensitivity and spectral overlap. The latter arises from the large number of protons resonat- ing in a rather limited proton chemical shift range (typically about 12 ppm for proteins). Matters however are rather different for the special case of paramagnetic proteins, which may show an increased proton spectral width, as well as characteristic relaxation properties, as described in Chapter 7 by Osborne et al, 1.2. A Simple Illustration Many of these characteristics of the NMR of proteins are illustrated by the simple example of the ‘H spectra of a moderately sized macromolecule, human transthyretin, a tetramer of four identical 127-residue subunits and an overall molecular weight of ca. 54,000. The bottom trace of Fig. 1 shows the spectrum of about 2.5 mg transthyretm in 0.5 mL 90% HzO/lO% D,O at 77°C (the molecule is very stable with respect to thermal denaturation). It is possible to assign some groups of resonances to particular proton types in the protein, for example secondary (backbone) and primary (sidechain) amide protons, aromatic sidechain protons, a-protons in regions of P-sheet structure and aliphatic methyl side-chain protons, some of which are shifted to high field, probably owing to ring current shielding effects from neighboring aromatic sidechams, as indi- 4 Reid et al Transthyretin in D20 nsthyretin in Hz0 I’~“11 Y II 7 6 5 4 3 ?. I ” Fig. 1. 500 MHz ‘H NMR spectrum of the protein transthyretm at pH 7 0, 77°C m 90% H,O/lO% D,O Protons m different chemical environments give rise to signals in different chemical shift ranges, as marked, the downfield region of the spectrum of the same protein about 8 h after lyophrlrzmg and redrssolutlon m pure D20, showmg the persistence of a large number of backbone amide protons protected from exchange with solvent deuterrum by the protein fold cated above the spectrum. Short T2 relaxation times have caused broadening of the resonances m the spectrum and obscured the spm-spm couplmgs. Many of the signals from the amide protons are still visible even tf the sample IS dts- solved m D20 (top trace of Fig. 1) instead of HZ0 because the fold of the protein occludes them from solvent and hence prevents or delays then exchange with deuterons. 2. Sample Considerations Most NMR expertments mvolvmg proteins are carried out m Hz0 (to which 5-10% by volume D20 has been added to provtde a field-frequency lock) or D,O. HZ0 has the advantage that peptrde backbone amide protons are visible, whereas m D20 they may disappear due to exchange with the solvent. Work- mg m HZ0 can create dynamic range problems for the analog-to-digital con- Introduction to the NMR of Proteins 5 verter (ADC) of the spectrometer because of the intensity of the water signal; however, numerous methods, ranging from simple presaturation to multipulse sequences designed for minimal excitation of water, to selective shaped pulses combined with pulsed magnetic field gradients, have been evolved to mini- mize the obtrusiveness of the water signal. NMR is an insensitive spectroscopic technique, and so any experiments aimed at total structure elucidation tend to be performed at concentrations of about 1 mM or above. Thus, limited solubil- ity, and aggregation, can limit the tractability of certain proteins. Because increasing the overall and local mobility of a macromolecule has the effect of decreasing resonance linewidth and increasing resolution, it is often desir- able to conduct protein NMR experiments at elevated temperatures. Obviously the protein must be stable at the temperature chosen. If either of these techniques is available, it may be helpful to check the thermal stability of an uncharacterized protein by circular dichroism (CD) or Fourier transform infrared (FT-IR) techniques (41), which are much less demanding of material, before risking large amounts of protein (or m&-unrent time!) on protracted NMR experiments. Many structure elucidation NMR experiments are carried out at acidic pH, between 3.0 and 5.0, because in weakly acid media the exchange rates of peptide amide protons with solvent water are at a mmimum, and they can be readily observed. Acettc and formic acids, which are available m deuterated forms, are suitable as buffers at these pHs if necessary. At higher pHs, around neutral, sodium or potassium phosphates are ideal as they do not contribute extra proton signals in the proton spectrum; however phosphate can cause aggregation or precipitation of some proteins, and deuterated “tris” (iris- hydroxymethylaminomethane, tris-d, i) is an acceptable alternative. When mea- suring acidity in D20, the actual deuteron ion activity, measured by the quantity pD, is related to a pH meter reading, pH*, by the relationship pD = pH* + 0.4. Freeze-drying (lyophilization) is an excellent method of bringing a dilute sample up to NMR concentrations, provided that the protein is stable to such treatment. However, nonvolatile cosolutes such as inorganic salts, buffers and detergents will be concentrated to the same degree as the protein during this procedure. Volatile buffers such as ammonium bicarbonate or ammonium acetate can for this reason be very useful in purification. Proteins which dena- ture on lyophilization can be concentrated by techniques such as ultrafiltration, and separated from cosolutes which will be spectroscopically troublesome by dialysis or size exclusion chromatography. Many proteins, especially those which associate with membrane or other lipids, can require solubilization with detergents like sodium dodecyl sulfate (SDS, commercially available from suppliers of stable-isotope labeled fine chemicals like Cambridge Isotope Labs, Woburn, MA, in deuterated form). Membrane-bound proteins cannot be studied by conventional high-resolution 6 Reid et al. solution NMR methods, and special techniques based on solid-state NMR are required, as described in Chapter 8 by Klassen and Opella. Nonaqueous solvents such as trifluoroethanol (TFE) and hexafluoroisopropanol (HFIP) are commonly used in peptide studies for their ability to solubilize many mem- branes associated and hydrophobic sequences at comparatively low ratios to water. These solvents also have structure-stabilizing properties so their use m the study of membrane and other peptides should be approached with caution, and Justified if possible by demonstrating correspondence between membrane- bound and solubilized conformations by a technique such as FT-IR (42). It is sometimes necessary to add reducing agents to protein solutions to pre- vent the spontaneous formation of unwanted intra- or intermolecular disulphide bonds. Mercaptoethanol and dithiothreitol are available m deuterated form and low (1 mM) concentrations usually suffice. The presence of even small (micro- molar) amounts of paramagnetic metal ions like manganese, nickel, copper, and iron can be mimical to high-resolution NMR and small amounts of EDTA (also commercially available deuterated) can be added to sequester them Sodium azide to a concentration of a few millimolar is often added to protein solutions to inhibit fungal and bacterial growth. It is often necessary to lyophilize a protein from one solvent, for example, H20, and reconstitute it in another such as D20 or TFE. This can often be done without removing the solution from the NMR tube by freezing it along the walls of the tube, the latter being cooled in, preferably, liquid nitrogen or acetone/dry ice. The same freezing technique can be used if it must be stored below OOC,b ecause samples simply allowed to freeze will expand and crack the NMR tube. 3. Fundamentals of Spectral Assignment 3.1. Sequence-Specific Assignment In order to define the three dimensional structure of a protein or determine how it interacts with ligands, it is first necessary to assign as many of its NMR resonances as possible to specific nuclei in individual amino acid residues. For small proteins of about seventy constituent amino acids, this can be done, essentially completely, by two-dimensional homonuclear spectroscopy and the sequence specific assignment procedure of Wtithrich and coworkers (43). This procedure has been well detailed in several text books, for instance by Redfield in ref. 27 and Neuhaus and Evans, and Williamson, in ref. 35, and is covered in detail in Chapter 11 by MacLachlan et al. Homonuclear, i.e., proton only, assignment methods start to break down when the complexity of the “fingerprint region” (containing amide NH to a-proton cross peaks) prevents unam- biguous assignment of crosspeaks to specific residues. As a rule this occurs for proteins of between about 50 and 100 residues (although it will be interesting Introduction to the NMR of Proteins 7 to see this ceiling raised by the dispersion offered by 750 MHz and higher field instruments). At this point, resonance assignment IS facilitated by rsotopic labeling of the protein-incorporation of NMR active heteronuclei, in particular 13C, 15N, and 2H-and the use of a range of newer experiments, as described in Chapter 2 by Whitehead and Waltho. The sequentral resonance assignment procedure will be illustrated by a simple example, based on the N- and C-terminally blocked disulphide-cychzed pentadecapeptide: which was synthesised as a potential “mimic” of a loop of the putative benzo- diazepine binding region of the GABA* receptor. The first step in any asstgnment procedure involves correlating NMR signals from protons that are J-coupled, and thus in effect separated from eachother by only two or three covalent bonds. This is done using variants of the COSY (Correlated Spectroscopy) and TOCSY (Total Correlation Spec- troscopy, also known as HOHAHA, for Homonuclear Hartmann-Hahn) experiments. These techniques enable thorough bond connectivities to be established between protons in the same amino acid residue. Figure 2A shows the fingerprint region of the double-quantum filtered (DQF) COSY spectrum of the peptrde, which corresponds to the crosspeaks due to the three bond connectivittes between the amide protons (the chemical shifts of which are plotted on the x, or F2, axis) and a-protons (chemical shifts plotted on they, or Fl, axis). As expected, a total of 13 cross peaks can be observed, since the prolmes do not give rise to a correlation in this region of the spectrum. It is often possible to identify glycine residues because they can give rise to two crosspeaks, NH-aH and NH-aH’. Although the fine structure of the COSY experiment does reflect the couplmg muliphcities of each mteractmg nucleus, accurate coupling constants cannot be measured directly from the spectrum; this can be done by spectral stmulations, or by using the Exclustve COSY experiment and its variants like the Primitive Exclusive COSY (PE-COSY). The TOCSY experiment enables correlations between protons separated from each other by several intervening protons to be observed, by varying the spmlock mixing time in the pulse sequence. This allows determination of the type of spin system (44) of which a resonance is part. Amino acids such as glycine, alanine, valine, leucine, isoleucine, and threonme give rise to unique patterns of crosspeaks in a long mixing time (rM of between 60 and 100 ms) TOCSY experiment. However, amino acids with AXY spin systems (NH-aH- PH, PH’), such as aspartic acid, asparagine, cysteine, phenylalanine, tyrosine, histidine, and tryptophan, and those with AMNXY spin systems (NH-aH- PHj3H’ -yH-yH’), such as glutamic acid, glutamine, and methionine, cannot be 8 Reid et al. A I 1 A14 1.0 1.2 I 1.2 -I 4 1.4 0 -1 6 tl 1.6 -1 8 P2 and we’s PZa nd P9ys :: 1.8 E c -2 0 0 07 5.0 -2 2 2.4 IPrn t PPrn 86 84 82 80 78 76 I 26 t B I t2.a ~L53.S -1 0 32 -1 5 38 -2 0 / pm -p m 4.4 4.2 4.0 3.8 3.6 3.4 25 / tm , 8.6 8.4 8.2 8.0 7.0 7.6 Fig. 2. Datasetss howing the determination of through-bond (J) connectivities in the cyclized GABA, receptor mimetic peptide in DMSO-d6. (A) Shows the fingerprint region of a DQF-COSY spectrumc ontainingt he crossp eaksa rising from coupling between the peptide amide protons and a-protons in eacha mino acid residue (except introduction to the NMR of Proteins 9 distinguished from each other m this way. The aromatic residues, phenylalanine, tyrosme, hlstldine, and tryptophan, can often be identified at a subsequent point by NOES between their aromatic protons, and /3- or a-protons of the same residue, and by characteristic chemical shifts of then aromattc protons. The side chains of argmme and lysine may frequently be identified by correlations in the long-range TOCSY spectrum from the side chain amino proton resonance and from the terminal guanidinium group. Prolines can often be Identified m the long-range TOCSY, m spite of lackmg NH protons, from the dtstmctive connecttvtties between a, /3$‘, y,y’ and 6,6’ protons Ftgure 2B shows the region of the long-range TOCSY spectrum portraying the connectivities arising from the NH protons and transmitted along the side chains of the ammo acids m the above sequence, while Ftg. 2C shows connectivities due to the side chains of the two proline residues in the sequence. Having assigned as many spm systems as possible to amino acid types, the next step in the sequential asstgnment procedure involves ascribing to each residue its specific place in the pepttde or protem sequence. Thts can be achieved by using the NOE experiment m, most commonly, its two-dlmen- slonal version, the NOESY (nuclear Overhauser effect correlated spectroscopy) or variant ROESY (rotating frame NOE correlated spectroscopy) The ROESY experiment may be useful for smaller peptrdes m which the molecular correlation time can result m the NOE being small or zero, although care should be taken with this experiment, since tt is possible to observe correlattons artsmg from sources such as homonuclear Hartmann-Hahn (HOHAHA) transfers, inter alia (39). The arm of sequential assignment IS to observe inter-residue NOES between residues that have been assigned previously to specific amino acid types. Some of the NOES commonly observed among peptide backbone protons are sketched below: the prolines and the N-terminus) (B) Shows how the long-range TOCSY expertment can delineated tfferent spin-systemsb y virtue of longer rangeJ -connecttvttresb etween peptrde amide protons and other protons m the same resrdue, while (C) shows the mamfestation of prolme spin-systemstn the samee xperrment. 10 Reid et al. mm mm 86 84 82 00 70 76 Fig. 3. A dataset rllustrating the NOE effects underlymg the sequence-specific assignment strategy for a fragment of the GABA receptor mrmetrc peptrde The figure shows the fingerprint region of a rotating frame NOE (ROESY) experiment acquired using a mixing time of 200 ms The arrows trace out the network of interresidue NOES; vertical arrows connect cross peaks corresponding to NOES between the same amide proton and different-probably adJacent-o-protons while horrzontal arrows connect srgnals from amide protons and the same a-protons. Figure 3 gives a simple illustration of how analysis of these sequential NOES leads to assignment. The lowest field amide proton signal m the spectrum has been assigned to a NH-aH-PH, PH’ spin system on the basis of the long-range TOCSY experiment. As expected, the amide proton gives an NOE to its own a-proton labeled with a “*,” which has been identified from the COSY and TOCSY, Fig. 2A and B), but it also gives an interrestdue NOE to another a-proton (signified by the vertical arrow tn Fig. 3 joining the first crosspeak to the one labeled with a “t”). There IS another cross peak at the same chemical shift as the o-proton of the interresidue cross peak (horizontal arrow to the crosspeak labeled with a “I#?‘), which corresponds to the chemical shift of the a-proton of one of the alanines in the sequence, identified by its unique long- range TOCSY connectrvity pattern. This is probably the intraresidue NH-aH cross peak for this alanine. The ammo acid sequence is then searched to find the number of possible occurrences of da&r + n) NOES (where n = 1,2,3) between an alanine residue and an amino acid giving rise to an AXY side chain