Transcript Lecture 10

Lecture 10: NMR Applications
We may know a tiny bit about NMR theory, but what
bioanalytical problems can we adress with NMR?
Organic Analysis: Specific organic molecules give
characteristic signals in terms of chemical shifts and peak
splitting due to J coupling
Solid state structures: NMR can be used on solids! But we
need to use a special trick, which we’ll talk about
Bioanalytical Applications
Protein Structures: NMR is one of two main ways of
acquiring molecular images of proteins
Protein/Protein, Protein/Ligand Interactions: NMR can be
used to ‘track down’ binding interfaces
Protein dynamics: NMR is the only way to get atomic level
information about how proteins move.
So You Want To Be An NMR Spectroscopist…
If you’re a bioanalytical NMR spectroscopist, here’s the
typical runup to an experiment:
1) Grow up your protein with the appropriate label. You’ll
either be expressing your protein in bacteria (probably E.
coli) or yeast (probably S. Cerevisiae)
For 2D NMR: Probably 15NH4Cl or 13C--D-Glucose
For 3D NMR: Probably 15NH4Cl and 13C--D-Glucose
2) Purify your labeled protein (probably His6 or GST tag)
3) Dilute to desired conc. (probably in water, around 1 – 10
mM), add 5-10% D2O, 100 uM DSS
4) Make sure the protein is stable under these conditions,
place in NMR tube, throw into instrument
So You Want To Be An NMR Spectroscopist…
5) Set the temperature, check tuning/matching
6) Shim the instrument: We have to make the magnetic field
perfectly homogenious across the sample or equivalent
nuclei will have different spins!
The computer can do this for you using the ‘gradient’ approach discussed
last time. Shims are extra magnetic coils with their fields pointed in
essentially every direction relative to the sample. They can therefore ‘add’
or ‘subtract’ to the big huge magnetic field as needed to even it out.
I
shim x
H(H2O)

shim z
So You Want To Be An NMR Spectroscopist…
7) Calibrate the hard pulse length for 90°:
z
time to
360°/4!!
z
-y

-y

8) You’re ready to go! Load and calibrate the experiment you
want to do!! You may have to work out the appropriate
power and/or duration of certain soft pulses, depending on
the experiment and water suppression scheme.
Good tutorial on biological nmr:
http://www.nmr.sinica.edu.tw/Cours/Course20040906/NMRExperiments_
LargerMolecule.pdf
More on Gradients: DOSY
We’ve said that you can destroy magnetization using a
gradient pulse. But you can also reconstitute it by using the
same pulse with the opposite phase at a later time:
Of course, this will only work if the nuclei remain essentially
stationary over the course of the wait period. This is the
basis for diffusion measurements (DOSY) by NMR!
Water Suppression
Biological experiments are carried out in water. If we want
to see protons from our sample we’re going to need to
strongly suppress the water signal.
Selective ‘soft’ pulses on
Here are a few ways of doing that:
H2O protons
Watergate:
1H
Defocus
everything
Gz
3-9-19
Watergate:
Defocus water,
refocus not water
Water Supression
Flipback Watergate:
Puts water on z before first gradient
pulse
Pre-saturation:
Lengthy, continuous ‘soft’ excitation
of water offset
NMR of Peptides
Now that we’ve suppressed our water signal, we can take
some spectra of peptides in water.
If we do a simple water suppression pulse-acquire experiment,
we may see something like this:
NMR of Peptides: TOCSY
That can be useful – we have some idea of what we’re
looking at… but which peak corresponds to which specific
proton?
TOCSY
TOCSY
tells us
which
amino acid
belongs to
which peak
NMR of Peptides: NOESY
But we still don’t know the amino acid sequence. For that
we need to look at ‘through space’ interactions:
NOEs are a relaxation effect.
As such they are dependent
on the correlation time:
NOE
1D Saturation Transfer
Saturation transfer is a simple technique that can be used to
determine if and how something small is binding to
something very big.
Saturation here is the same as presaturation in water suppression. It
involves continually hitting a select
frequency with a train of soft pulses:
Sat. pulse
Protein NMR: We Need More Nuclei
1D and H-H coupling experiments are all well and good
when you’ve got < 200 protons, but proton signals are not
really all that well dispersed.
We’re going to need to use couplings between two different
nuclei (heteronuclear NMR). Since we’re dealing with
proteins our options are most likely 13C (very expensive!)
and 15N (expensive).
The most dispersed signals would be the carbons, but
nitrogen is much cheaper!
Thus, the most common type of protein NMR spectrum is
an HSQC which usually correlates the amide nitrogen with
the amide proton. Thus there is one peak per residue (except
proline!!)
A 13C Protein Spectrum
Here’s a 13C Spectrum of an SH3 domain (aprox. 70 a.a.):
HSQC Spectra
Here’s the pulse sequence for the HSQC experiment:
y
I
x/-x
S
And here’s the result:
This is ‘the’ HSQC
for properly folded
Sso Acylphosphatase
(104 a.a.)
Detecting and Localizing Ligand Binding
Most analytical techniques work hard to tell us ‘if’
something is binding to our protein of interest. NMR not
only tells us that, but where!
The most common way of measuring this is by ‘ligand
titration’ experiments which amount to monitoring the
HSQC as a function of ligand concentration.
Low Ligand
High Ligand
Med Ligand
Protein Dynamics by NMR: H/D Exchange
Once you’ve got an HSQC, you can study slow (minutes to
days) conformational dynamics by NMR.
To do this, you calibrate your HSQC for speed, ‘buffer
exchange’ your protein into 90%+ D2O, RUN to the NMR
instrument, drop your sample in, quick re-shim and GO!!
Here’s the result:
1st HSQC after D2O
No D2O
t = 60 min
HDX results
Since we know which backbone protons correspond to
which signals, we can identify which are more protected:
H/H Exchange: CLEANEX
CLEANEX is a cool HD exchange technique that uses water
protons instead of 2H!
Backbone and water protons are exchanging all the time
Instead of exciting all protons except water, we only excite
water
These magnetized protons now exchange onto the protein…
And we use that magnetization to transfer to 15N
H
H
H
O
Results of CLEANEX
In order for CLEANEX to work, exchange has to occur faster
than the relaxation of protons on the protein. This means
mid-to-low milliseconds range:
Sequencing Proteins by NMR
The HSQC gives us a spectrum in which each amino acid is
distinguishable, but doesn’t tell us much about which amino
acid they are, and in what order. To do that, we need to
extend our analysis into the 13C plane. 3D NMR!!
Sequential Assignment by NMR
To do ‘sequential assignments’, we use pairs of J-couplingbased 3D experiments, the most common pair is:
HNCA
C1
N
C
H
O
H
O
N
C
C2
HNCOCA
C1
N
C
H
O
H
O
N
C
C2
Getting Structural Info: The CSI
The Chemical Shift Index (CSI) is a quick way of assessing
secondary structure:
RESIDUE TYPE HA
CA
CB
CO
To your observed shifts, give score:
+1 if >.7 ppm higher than CSI value
-1 if >.7 ppm lower than CSI value
0 if within -.7 to +.7 of CSI value
Four shifts in a row at -1 HA and/or
+1 CA/CO = minimum for Helix
Three shifts in a row at +1 HA and/or
-1 CA/CO = minimum for -strand
All other regions are designated
random coil
Ala
4.35
52.5
19.0
177.1
Cys
4.65
58.8
28.6
174.8
Asp
4.76
54.1
40.8
177.2
Glu
4.29
56.7
29.7
176.1
Phe
4.66
57.9
39.3
175.8
Gly
3.97
45.0
-
173.6
His
4.63
55.8
32.0
175.1
Ile
3.95
62.6
37.5
176.8
Lys
4.36
56.7
32.3
176.5
Leu
4.17
55.7
41.9
177.1
Met
4.52
56.6
32.8
175.5
Asn
4.75
53.6
39.0
175.5
Pro
4.44
62.9
31.7
176.0
Gln
4.37
56.2
30.1
176.3
Arg
4.38
56.3
30.3
176.5
Ser
4.50
58.3
62.7
173.7
Thr
4.35
63.1
68.1
175.2
Val
3.95
63.0
31.7
177.1
Trp
4.70
57.8
28.3
175.8
Tyr
4.60
58.6
38.7
175.7
Getting Structural Info: NOEs
In NMR, the Nuclear Overhauser Effect is the effect that one
nucleus has on the relaxation of another. The intensity of
this effect is directly related to the proximity of the
interacting nuclei:
1
NOE 6 f ( c )
r
‘is proportional to’
absolute distance between
the interacting nuclei
correlation function – describes
attenuation (or buildup) of the NOE due
to the relative motions of the nuclei
So the internuclear distance effects the size of the NOE
Just like in J coupling, NOE coupled nuclei will experience
an oscillating phase at each other’s offsets .
This tells us which nucleus is interacting with which, but a 3D
experiment (e.g. HSQC-NOE) is required to distinguish
THE HSQC-NOE experiment
Here’s the most common NOE-based experiment for
structure elucidation:
Has the advantage of not requiring
double labeling
Gives us a set of inter-proton
distance contraints
We know which amide proton is
which and which amide protons
are nearby (1.6 – 6 Å).
More Structural Info: Angle Restraints
A network of NOEs from an HSQC-NOE is a start, but
there’s plenty it doesn’t tell us, particularly which way the
side chains are pointed.
One ‘cheating’ way is to use / values consistent with the
secondary structures derived from CSI, but there are weak
constraints
A better way is to do a ‘residual dipolar coupling’ experiment
in which the sample is placed in a medium, such as
polyacrylamide or phage coat particles, that causes a net
alignment with the magnetic field.
B0
Residual Dipolar Couplings
Through bond (J) dipolar couplings have well defined
frequencies called coupling constants. It is, in fact, by using
coupling constants that we pass magnetization around
through bonds (such as in a TOCSY).
J1,2
J1,2
The magnitude of the coupling constant
depends on the orientation of the interaction
with respect to the big huge magnetic field.
I

By measuring the coupling constant (which
can only be done in alignment media), we can
figure out the bond angle with respect to B0.
difference between
aligned and
unaligned J
angle with respect
to B0
Results of Residual Dipolar Couplings
Here’s what RDC’s look like. We have to
run our HSQC without decoupling.
Now we have distance constraints and some
bond angles. Combined, these are sufficient
to allow us to parameterize a model protein
structure.
The next step is to plug our distance and
angular constraints into a computer program
that uses a molecular mechanics force field to
find the lowest energy structures that meet
the constraints.
NMR Structure Results
Here’s what an NMR ‘structural ensemble’ looks like:
You can then take the average structure
or the ‘best’ structure (the one that best
fits the constraints) to give you a final
structure:
Note that, unlike x-ray crystallography,
this is a structure from the protein in
solution.
There are currently 4,448 protein
structures in the BioMagResBank database.
Dynamics by NMR
Since we’re looking at our protein in solution, it should also
be moving around roughly like it does in vivo. NMR will
allow us to get site specific information about these
movements.
NMR is also the only method by which motions on virtually
all relevant time-scales are observable:
Virtually every type of
protein motion/activity is
covered.
Very Fast Motions: T1 and T2
We talked about longitudinal (T1) and transverse (T2)
relaxation (biophysicists call them R1 and R2)
To make a very long story short, you can get a general
description of the conformational freedom for nuclei in a
protein by mapping out the spectral density function J()
which is directly related to T1 and T2.
The spectral density function at any particular frequency 
is related to the order parameter S via the following
m = correlation time
relationship:
 = correlation time + timescale of
bond vibration (~ns)
An order parameter of 1 indicates complete restriction of fast
timescale motions while S = 0 indicates completely
unrestricted motion.
Slower Motions: Relaxation Dispersion NMR
Until recently, there was a big hole in the timescale
accessible to NMR measurements. And it was centered right
on the all important millisecond timescale.
The reason is that the equation for the spectral density function
becomes underdetermined when an
additional term to account for
conformational exchange is added.
The answer was developed at nearby U of T in the Kay group.
They advanced a technique called Carr-Purcell-Meiboom-Gill
(CPMG) Relaxation Dispersion NMR.
CPMG relaxation dispersion
The key to CPMG relaxation dispersion is that the
contribution to J() from Rex is suppressed by the
application of a train of  pulses.
As a consequence, the
contribution to J() from Rex
alone can be measured and the
frequency of the motion causing
Rex can be acquired.
A major advantage of CPMG RD
is that is sensitive even to low
population protein folding
intermediates.
Chem. Rev. 2006, 106, 3055-3079
Solid State NMR
Remember we said we can’t see big stuff in solution by
NMR because to correlation time is too long and thus T2 is
too fast. Well, what about solids?
They aren’t tumbling at all, so they have infinitely long c and
thus an (almost) infinitely fast T2. (Also recall that, after an
initial rise, T1 goes down with increasing c).
BUT – this T2 relaxation is almost all secular, meaning that it
is due primarily to dipolar couplings
In solution, random tumbling causes these dipolar couplings,
which are vector quantities, to cancel each other out
Fortunately, some very clever people thought of another way
of making this happen…
Solid State NMR
In solid state NMR, we tilt the sample to the ‘magic’ angle,
which is 54.74° relative to B0.
B0
=54.74°
And then we spin it around that angle at very high
frequency. Thus the name of this type of NMR – ‘Magic
Angle Spinning’.
To be effective, this spinning has to be close to or above the
offset frequency  of the nuclei being observed.
We don’t use this too much in bioanalytical chemistry… YET!
We’re Done!!!
At times, you might have
felt like this…
But now you’re almost
like THIS!