Protein NMR Techniques

410 Pages·1997·27.055 MB·English

by  David G. Reid

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 sec-
ondary 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), for-
mally 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 correla-
tion 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 paramag-
netic 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 mol-
ecule 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 side-
chain 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) tech-
niques (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 sol-
vents 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 “finger-
print 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 particu-
lar 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 sig-
nals 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 finger-
print 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, phenylala-
nine, tyrosme, hlstldine, and tryptophan, can often be identified at a subse-
quent point by NOES between their aromatic protons, and /3- or a-protons of
the same residue, and by characteristic chemical shifts of then aromattc pro-
tons. 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 con-
nectivities 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 correla-
tion 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 pro-
tons 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