Peptide Bond Geometry Studied by Solid-State NMR Spectroscopy
THESIS
Presented in Paritual Fulfillment of the Requirements for the Degree of Master of Science
in the Graduate School of The Ohio State University
By
Chitrak Gupta,
Graduate Program in Chemistry
The Ohio State University
2013
Master’s examination committee:
Professor Christopher Jaroniec, Advisor
Professor Thomas Magliery
Copyright by
Chitrak Gupta
2013
Abstract
Three-dimensional structure of proteins is an intrinsic property of every protein,
and is directly related to its biological function. Studying protein structure is thus of
immense importance to understand the mechanism by which the proteins perform such
functions. In principle, the backbone structure of a protein can be completely described
by a set of torsional angles. Indeed, a significant amount of structural studies of proteins
have involved measuring the torsional angles defined by C’-N-Cα-C’ atoms (denoted φ)
and N-Cα-C’-N atoms (denoted ψ). Little has been done, however, to study the third of
the triplet, the ω-torsional angle, defined by the Cα-C’-N-Cα atoms, which is usually
assumed to be planar, with the atoms arranged in a trans-like conformation. However,
cis-like peptide bonds are known to exist, often at or near active sites, which make them
biologically significant, although there are no reliable experimental methods apart from
crystallographic studies to differentiate cis-peptide bonds form their trans- counterparts.
This thesis is aimed at developing a new solid-state NMR experiment to study the
peptide bond geometry. The primary objective is to differentiate cis- and trans-peptide
bonds in polypeptides and proteins. Chapter 2 describes the synthesis of isotopically
labeled model compounds with trans- and cis-peptide bond. The former, glycylglycine,
was synthesized by solid phase peptide synthesis following standard protocols, with
minor modifications to increase the yield. The latter, 2,5-diketopiperazine, was
ii
synthesized using microwave-assisted synthesis, a protocol recently published. Both the
compounds could be obtained in high purity and integrity, as shown by their solution 1H
and 13C NMR spectra.
Chapter 3 describes the design of the NMR pulse sequence which can achieve this
goal. The fundamental idea is to allow correlated evolution of two different anisotropic
interactions under magic-angle spinning. Such evolutions are sensitive to the relative
orientation of the two interactions. In this case,
13 α 15
C - N dipolar coupling and
13
C’-13Cα
dipolar coupling have been correlated, to distinguish between the two geometries about
the N-C’ bond. The possibility of using
13
13
C’ chemical shift anisotropy instead of
13
C’-
Cα dipolar coupling has been discussed. Preliminary simulations have shown the
detection of a cis-peptide bond to be possible, although, thus far, the experiment is not
sensitive enough to measure deviation from planarity of a peptide bond.
Chapter 4 shows the solid-state NMR spectra obtained from these model
compounds. The 1D
13
C and
15
N CP-MAS spectra from both the compounds confirm
their purity and integrity, and provide sufficient signal-to-noise to proceed with the
proposed experiment. REDOR dephasing recorded on both the compounds are in good
agreement with simulation. Chapter 5 discusses the future perspectives of this work,
including extending it to bigger systems, and to use
iii
15
N-1H dipolar coupling instead of
13
C’-13Cα dipolar coupling to increase sensitivity and enable measurement of the ω-
torsional angle with reasonable accuracy.
iv
Dedication
Dedicated to all hard workers awaiting recognition
v
Thank you Ma, Baba, and Ananya.
vi
Acknowledgments
I shall take this opportunity to thank the variety of people without whose help this
thesis would not have been a success. First of all, I would like to thank my advisor, Dr.
Christopher Jaroniec, for his extremely helpful guidance throughout the course of my
study in his research group. I would also like to thank all the former and current members
of the group for the wonderful experience I have had working with them.
I specially thank Judith Brown, Graduate Program Coordinator, and Jennifer
Hambach, Graduate Admissions Coordinator, at the Department of Chemistry of The
Ohio State University. Before and immediately after my arrival at OSU, I was full of
confusions regarding the way the administrative processes works in this country, and
Jennifer was always ready with help. Throughout my stay in the program, I have been
immensely helped by the timely reminders from Judy, not to mention the multiple times I
have been at her office with the strangest of problems. One of the most approachable
people I have ever known, Judy has always had the answers to my questions.
I would thank my parents for their unconditional love and support throughout my
life, and for always encouraging me to pursue my dreams. I thank them for teaching me
that life is bigger than success and failure, which has given me the strength to go on in
difficult times. I would extend my gratitude to my high school teachers, speficially, Dr.
vii
Sekhar Pal, for getting me interested in Chemistry. I am grateful for having been taught
by encouraging teachers, both at St. Stephen’s College and IIT Roorkee. My teachers not
only helped me to grasp a fundamental understanding of Chemistry, but also inspired me
to aim high.
I have been extremely fortunate to have friends, both in Columbus and outside,
who has made life livable. I would thank Souvagya, Chiranjit, Dwaipayan, Mithila, Arijit
and Shiladitya for making my stay away from home a lot easier. I would forever cherish
the potlucks and movie-sessions with Arijit and Shiladitya. I would never learn to cook
and to make an apartment look decent, without Chiranjit. I am indebted to Mithila for
teaching me to play squash and getting me to play table-tennis after fifteen years. No
amount of gratitude is sufficient for Souvagya, who I could always bump into, for
anything good and bad, but most importantly to discuss plans for long trips. My
knowledge of history and digital electronics (and pretty much everything else) has
increased significantly since meeting Dwaipayan. Some of the best times of my life have
been those spent with each one of you at Buckeye Donuts, the coffee shop that by itself
deserves special mention. I would thank Shreya and Joyeeta for always being just a
phone call away, keeping me connected to the life I left behind in India.
Last but not the least, I would express very special gratitude to Ananya, for being
by my side and continuing giving me hope through thick and thin. I can never thank you
enough for keeping me cheerful even during my darkest times.
viii
Vita
April 22, 1986……………………….………………………………Born, Kolkata, India
2004………………………………………..Salt Lake School (Eng. Med.), Kolkata, India
2007………………………………..……………..B.Sc., University of Delhi, Delhi, India
2009………………..……M.Sc., Indian Institute of Technology Roorkee, Roorkee, India
2009-present………………………………..Graduate Teaching and Research assistant,
Department of Chemistry, The Ohio State University
Fields of study
Major Field: Chemistry
ix
Table of contents
Abstract…………………………………………………………………………ii
Dedication……………………………………………………………………....v
Acknowledgement……………………………………………………………...vii
Vita………………………………………………………………………………ix
List of tables……………………………………………………………………..xiii
List of figures……………………………………………………………………xiv
Abbreviations……………………………………………………………………xxi
Chapter1:Introduction………………………………………………………………..1
1.1: Peptide bond geometry…..........................................................................1
1.1.1: Planarity of peptide bonds…………………….…………….…1
1.1.2: Unusual peptide bonds in proteins…………………………….2
1.1.3: Biological significance of non-proline cis-peptide bonds….…5
1.1.4: Proposed methodology: A solid-state NMR approach………..6
1.2: NMR spectroscopy……………………………………………………...8
1.2.1: Matter and spin………………………………………………..8
1.2.2: Zeeman splitting………………………………………………8
1.2.3: Fourier-transform (FT) NMR………………………………...11
x
1.2.4: Multidimensional NMR…………………………….………..12
1.2.5: Magnetic interactions……………………………….……….13
1.2.6: Solid-state NMR and magic-angle spinning………….…......20
1.2.7: Recoupling experiments……………………………….…….21
1.2.8: Measuring torsional angles………………………….………22
Chapter 2: Synthesis of model compounds……………………………………….24
2.1: Synthesis of model compound with trans-peptide…………………...25
2.1.1: Problems associated with solid phase peptide synthesis…...43
2.1.2: Synthesis of (U-13C2, 15N) glycylglycine…………………..44
2.2: Synthesis of model compound with cis-peptide……………………..47
2.2.1: Synthesis of 2,5-diketopiperazine with solid phase peptide
synthesis…………………………………………………….……47
2.2.2: Synthesis of 2,5-diketopiperazine with microwave-assisted
synthesis………………………………………………….………48
2.2.3: Synthesis of of (U-13C2, 15N) 2,5-diketopiperazine……….53
Chapter 3: Design of NMR experiment………………………………………….55
3.1: Recoupling experiments…………………………………………….55
xi
3.1.1: Heteronuclear dipolar recoupling…………………….....55
3.1.2: Homonuclear dipolar recoupling……………………….58
3.1.3: Recoupling of chemical shift anisotropy………………60
3.2: Cα-N-C-Cα experiment…………………………………………...60
3.2.1: Choice of C-C recoupling technique…………………..62
3.2.2: Choice of ratio of mixing times………………………..64
3.3: Utilizing carbonyl CSA for ω-angle measurement………………68
Chapter 4: Preliminary results……………………………………………….73
4.1: Solid-state 1D CP-MAS experiments…………………………...73
4.2: REDOR experiments……………………………………………77
Chapter 5: Conclusions and future work……………………………………80
References……………………………………………………………….….83
xii
List of tables
Table 1.1: Fraction of Xaa-Pro and Xaa-nonPro cis peptides as a function of structural
resolution………………………………..……………………..…4
xiii
List of figures
Figure 1.1: Frequency distribution of ω-angle, energy difference from planar structure in
peptides and proteins …………………..……………………………...…………1
Figure 1.2: Trans and cis peptide bond with and without proline…......................2
Figure 2.1: Glycylglycine…………..……………………………………………24
Figure 2.2: 2,5-diketopiperazine……….………………………………………..24
Figure 2.3: Proposed scheme for synthesis of glycylglycine…............................25
Figure 2.4: 1H NMR of glycylglycine (Bachem)…………….………………….26
Figure 2.5: 13C NMR of glycylglycine (Bachem)…………………………….…27
Figure 2.6: 1H NMR of product obtained by proposed scheme………..…….….29
Figure 2.7: 13C NMR of product obtained by proposed scheme...........................29
xiv
Figure 2.8: 1H NMR of Fmoc-protected glycine (AAPPTec) in CDCl3 with TMS
standard...............................................................................................................31
Figure 2.9: 1H NMR of Fmoc-protected glycine after attachment to resin and cleavage,
dissolved in CDCl3 with TMS standard………………………….……………..31
Figure 2.10: 1H NMR of product obtained by treating resin-bound Fmoc-Gly-Gly with
50% TFA in DCM, dissolved in CDCl3 with TMS standard...………………….32
Figure 2.11: 1H NMR of product obtained after cleavage in DCM……………...34
Figure 2.12: ESI mass spectra of product obtained after cleavage in DCM……..35
Figure 2.13: 1H NMR of product obtained with modified scheme……………….36
Figure 2.14: 13C NMR of product obtained with modified scheme………………36
Figure 2.15: ESI mass spectra of product obtained with modified scheme………37
xv
Figure 2.16: Overlaid 1H NMR of product obtained with modified scheme, with Fmoc
removal done for 30 minutes, 4 hours and 24 hours…………………….……..39
Figure 2.17:
13
C NMR of product obtained with modified scheme, with Fmoc removal
done for 24 hours................................................................................................40
Figure 2.18: The optimized synthesis scheme that was followed for the synthesis of (U13
C2, 15N) glycylglycine…………………………………………………….…41
Figure 2.19: RP-HPLC of glycylglycine at 220 nm……………………...…....42
Figure 2.20: Overlaid 1H NMR of glycylglycine (Bachem) and glycylglycine prepared by
the optimized scheme………………………………………..…………….…42
Figure 2.21: HCl crystals of glycylglycine…………………………………...43
Figure 2.22: 1H NMR of (U-13C2, 15N) glycylglycine……………………….46
Figure 2.23: HCl crystal of (U-13C2, 15N) glycylglycine……………………..46
xvi
Figure 2.24: Overlaid
13
C NMR of product obtained by microwave-assisted synthesis of
2,5-diketopiperazine, with hold times of 3, 6 and 9 minutes….……………...49
Figure 2.25: Overlaid
13
C NMR of product obtained by microwave-assisted synthesis of
2,5-diketopiperazine, with hold times of 15, 30 and 45 minutes…………….49
Figure 2.26: Overlaid 13C NMR of 2,5-diketopiperazine (Sigma) and 2,5-diketopiperazine
synthesized by microwave-assisted synthesis……………………………….50
Figure 2.27:
1
H NMR of 2,5-diketopiperazine synthesized by microwave-assisted
synthesis…………………………………………………………………...….50
Figure
2.28:
Reaction
profile
for
microwave-assisted
synthesis
of
2,5-
diketopiperazine…………………………………………………………..……51
Figure 2.29: Overlaid HPLC at 220 nm of 2,5-diketopiperazine (Sigma) and 2,5diketopiperazine synthesized by microwave-assisted synthesis………..……..52
Figure 2.30: Overlaid
synthesized
by
1
NMR of 2,5-diketopiperazine (Sigma) and 2,5-diketopiperazine
microwave-assisted
synthesis
after
purification…………………………………………………………….………52
xvii
HPLC
Figure 2.31: Reaction profile for the microwave-assisted synthesis of (U-13C2,
15
N) 2,5-
diketopiperazine………………………………………………….............………53
Figure 2.32: 13C NMR of (U-13C2, 15N) 2,5-diketopiperazine……………………54
Figure 3.1: Pulse sequence for Rotational Echo Double Resonance (REDOR)
experiment………………………………………………………………………..56
Figure 3.2: Simulated 13C-15N REDOR dephasing………………………………57
Figure 3.3: 15N-1H dephasing pattern..…………………………..……………….57
Figure 3.4: Schematic representation of the pulse sequence for measuring relative
orientation of αC-N and C’- αC internuclear vector………………………………61
Figure 3.5: Schematic representation for the same experiment with increased
dimensionality……………………………………………………………………62
Figure 3.6: Simulations for the choice of
13
C-13C recoupling element. a. DRAWS b.
MELODRAMA c. SPC-5 d. POST-C7…………………………………….......64
xviii
Figure 3.7: Simulated dephasing spectra of REDOR and POST-C7…………..65
Figure 3.8: Simulations for the choice of ratio of REDOR and POST-C7 mixing time. a.
REDOR mixing time = POST-C7 mixing time. b. REDOR mixing time = 2 * POST-C7
mixing
time.
c.
REDOR
mixing
time
=
0.5
*
POST-C7
mixing
time……………………………………………………………………………..67
Figure
3.9:
Sensitivity of simulated
dephasing
to
variation of ω-torsional
angle…………………………………………………………………………….68
Figure 3.10: Schematic representation of the pulse sequence for measuring the relative
orientation of the αC-N bond with carbonyl CSA……………………...............69
Figure 3.11: Simulated decay curve for the ROCSA experiment……………...71
Figure 3.12: Schematic representation of the pulse sequence for measuring the relative
orientation of 15N-1H internuclear vector and carbonyl CSA…………………72
Figure 4.1: 1D
13
C CP-MAS spectrum of (U-13C2,
15
N) glycylglycine at magnetic field
strength 500 MHz and 11.904 kHz magic-angle-spinning…………..………..74
xix
Figure 4.2: 1D
15
N CP-MAS spectrum of (U-13C2,
15
N) glycylglycine at magnetic field
strength 500 MHz and 11.904 kHz magic-angle-spinning…………..………75
Figure 4.3: 1D 13C CP-MAS spectrum of (U-13C2, 15N) 2,5-diketopiperazine at magnetic
field strength 500 MHz and 11.904 kHz magic-angle-spinning…………….75
Figure 4.4: 1D 15N CP-MAS spectrum of (U-13C2, 15N) 2,5-diketopiperazine at magnetic
field strength 500 MHz and 11.904 kHz magic-angle-spinning……………..76
Figure 4.5: 1D 13C spectrum of glycylglycine after CP transfers to
13
15
N and selectively to
C (carbonyl)…………………………………..………………...................77
Figure
4.6:
Experimental
and
simulated
REDOR
dephasing
pattern
of
glycylglycine…………………………………………………………………78
Figure 4.7: Experimental and simulated REDOR dephasing pattern of 2,5diketopiperazine………………………………………………………………79
xx
Abbreviations
C’
Carbonyl carbon
CP
Cross Polarization
CP-MAS
Cross Polarization Magic Angle Spinning
CSA
Chemical Shift Anisotropy
DIC
N,N’-Diisopropylcarbodiimide
DCM
Dichloromethane
DMAP
4-Dimethylaminopyrdine
DMF
Dimethylformamide
DRAMA
Dipolar Recovery At the Magic Angle
DRAWS
Dipolar Recovery with A Windowless Sequence
ESI
Electrospray Ionization
Fmoc
Fluorenylmethyloxycarbonyl
FID
Free Induction Decay
FT
Fourier Transform
FT-NMR
Fourier Transform Nuclear Magnetic Resonance
HPLC
High Performance Liquid Chromatography
HOBt
Hydroxybenzotriazole
MAS
Magic Angle Spinning
xxi
MELODRAMA
MELding Of spin-locking and DRAMA
NMR
Nuclear Magnetic Resonance
POST-C7
Permutationally Offset-STabilized C7
REDOR
Rotational Echo Double Resonance
RP-HPLC
Reversed Phase High Performance Liquid Chromatography
ROCSA
Recoupling Of Chemical Shift Anisotropy
SPC-5
Supercycled Permutationally Offset-STabilized C-5
TFA
Trifluoroacetic Acid
T-MREV
Transverse MREV
TMS
Tetramethylsilane
xxii
Chapter 1
Introduction
1.1
Peptide bond geometry
1.1.1 Planarity of peptide bonds
One of the most fundamental assumptions in protein structure is the assumption of
peptide bonds being planar. This is due to partial double bond character of the N-C’
bond. This restricts peptide bonds to two possible conformations around N-C’ bond,
where the ω torsional angle has value of 180º (trans) and 0º (cis) respectively. This was
described by Corey and Pauling1 in1953 where they noted that (i) N-C’ bond has ~40%
double bond character, and (ii) planarity of peptide bonds is a “sound structural
principle”.
However,
in
1968,
Ramachandran2
recognized
the
need
for
including
the
possibility of cis
Figure 1.1: Figure taken from reference 6. Frequency distribution of ω angle
(histogram), energy difference from planar structure for peptide (gray) and
protein (black), from reference 5. Red points are the similar energies calculated
from the distribution shown in histogram.
1
peptides in
structural
studies. The same group later showed3 that for peptides with bulky side chains,
agreement between measured and calculated 3JHN-Hα can be improved by considering nonplanar peptide bonds. Much later, in 1991, Pople’s group used high level Hartree-Fock ab
initio calculations to show4 that peptide bonds show significant flexibility about the N-C’
bond in the gas phase, with little energetic cost. Five years later, MacArthur and
Thornton surveyed5 peptide and protein databases to show that experimentally derived
structures show significant deviation from planar peptide bonds. They also calculated
peptide bond energy wells around ω = 180º from ω angle distributions using MaxwellBoltzmann relationship, which were further updated by Edison6. Figure 1.1, taken from
reference 6, shows the frequency distribution of ω angles along with original (reference
5) and updated values for energy of deviation of peptide bonds about planarity.
1.1.2 Unusual peptide bonds in proteins
In addition to assumption of
planarity, peptide bonds are usually
assumed
to
be
in
the
trans
conformation. Indeed, most peptide
bonds
exist
conformation.
in
This
the
can
trans
be
explained qualitatively by the fact
that cis-peptides have significantly
higher energy thant their transFigure 1. 2: Figure taken from reference 7. Trans and cis
peptide bonds with and without Proline
2
counterparts.
Energy
difference
between trans- and cis-peptide bonds is ~ 10 kJ/mol for peptides in which C-terminal
residue is anything other than Proline (Xaa-nonPro). This difference is ~ 2 kJ/mol if the
C-teriminal residue is Proline (Xaa-Pro). This makes trans-peptides far more abundant
than cis for both the categories.
However, an attempt to quantify the above argument brings up a discrepancy.
Based on these energy values, ~30% of C-terminus proline-containing (Xaa-Pro) peptides
and ~1.5% of C-terminus non-proline (Xaa-nonPro) peptides should exist in the cis
conformation. However, the fraction of peptides observed to exist in the cis conformation
is significantly lower8, both for proline and non-proline peptides. Even more intriguing is
the fact that the observed discrepancies are quite difference for proline and non-proline
peptides. While observed cis-peptide bonds are lower than expected by a factor of ~ 6 for
Xaa-Pro, the same is lower by a factor of ~ 50 for Xaa-nonPro. It is interesting to note
that high resolution (< 2.0 Å) structures contain almost twice the number of cis Xaa-Pro
bonds and four times the number of cis Xaa-nonPro bonds than medium and low
resolution (> 2.5 Å) structures. This has been tabulated by Weiss and Hilgenfeld 8 (Table
1.1).
This led the authors to believe that all the cis-peptides have not been identified. The
authors also noted that most structure refinement programs allow for the possibility of a
cis-peptide only in cases of Xaa-Pro unless specified explicitly. This immediately
underlines the importance of direct measurement of cis-peptides, especially for XaanonPro, through experimental methods. Also to be noted is the observation made by
Weiss and Hilgenfeld9 that in a structure where a cis-peptide is the correct structure, a
3
trans- conformation may be modeled with only the amide Nitrogen atom being displaced
by 1.0 Å. In totality, there is a definite need to experimentally determine cis-peptide
Table 1.1: Taken from reference 8. Fraction of Xaa-Pro and Xaa-nonPro cis peptides as a function of
structure resolution
< 2.0 Å
2.0 Å – 2.5 Å
≥ 2.5 Å
of 571
291
184
96
of 153209
72576
52194
28448
All
Number
proteins
Number
peptide bonds
Xaa-Pro
7413
3407
2566
1440
Xaa-nonPro
145796
69160
49628
27008
232 (0.32%)
140 (0.27%)
55 (0.19%)
Number of cis 427 (0.28%)
peptide bonds
Xaa-Pro
386 (5.21%)
205 (6.02%)
129 (5.03%)
52 (3.61%)
Xaa-nonPro
41 (0.028%)
27 (0.039%)
11 (0.022%) 3
(0.011%)
bonds. While NMR spectroscopy can be used to distinguish Xaa-Pro trans- and cispeptides10-12, till date, no general methods exist for this purpose, apart from X-ray
crystallography. The motivation, thus, is to develop a new experimental technique, that
will reliably differentiate cis- and trans-peptide bonds, without depending on the presence
of Proline at the C-terminus of the peptide.
4
1.1.3 Biological significance of non-proline cis-peptide bonds
Although extremely low in frequency, non-proline cis-peptides often have an impact on
the biological function of the protein they are a part of. Herzberg and Moult 13 examined 7
non-proline cis-peptide bonds, 6 of which were involved in ligand binding and/or
catalytic activity. Later on, Weiss et. al. 14 reported a refined structure of cellular factor
XIII, where they observed two non-proline cis peptides. As discussed by the authors, one
of these contain the active site at its N-terminus, while the other is close to the
dimerization interface. The authors also noted that the lack of any geometric need for a
cis-peptide at either of these locations. They concluded that cis-trans isomerism of these
peptides was the trigger for a conformational rearrangement, which provides a part of the
energy for binding to the substrate. The same year, Stoddard and Pietrokovski15 reported
the role of a non-proline cis-peptide in the self-splicing of DNA gyrase A. This and other
instances of strained amino acid geometries in binding sites and catalytic locations
instigated the authors to conclude that such geometries have “stored potential energy that
may be used to drive biochemical reactions and other physical processes in the cell”.
Non-proline cis-peptides have also been implicated in amyloid fibril formation,
although a conclusive picture has not been reached as yet. Spencer et. al. 16 used solidstate NMR spectroscopy to measure 13C-13C distances in a 9-residue peptide representing
the C-terminus of amyloid beta (Aβ), a major constituent of amyloid plaques that
Alzheimer’s Disease and other related dieseases. From their results, they concluded that
Gly37-Gly38 segment of β34-42 exists in cis-conformation. This result has been disputed
later on17 by the same group by taking into account -inhomogeneous broadening effects
that tend to make
13
C-13C distances longer than they actually are. However, even in this
5
work they could not confirm whether the said peptide bond is cis or trans, and only
concluded that both possibilities were likely.
Above examples highlight the need for an experimental technique capable of
distinguishing trans- and cis- peptide bonds. The technique needs to be general, i.e., be
able to pick up cis-peptides involving any two residues. Such a development, we hope,
shall enable refinement of protein structure databases in terms of frequency of cispeptides, especially ones not containing a Proline residue at the C-terminus. Given the
apparent role of such structurally-strained peptides in biological function, such a
technique could potentially throw light on the mechanism of functioning of a variety of
proteins.
In addition, we tried to take this a step further. We wanted to see if, in addition to
distinguishing between trans and cis peptide bonds, we could measure the deviation from
planarity of a peptide (section 1.1.1). Depending on the sensitivity of this new
experiment, it might be possible to obtain observable difference in the behavior of
peptide bonds of varying ω angles. This will be discussed in Chapter 5.
1.1.4 Proposed methodology: A solid-state NMR approach
With the problem defined and the importance outlined, proposed methodology for
attacking the same shall be described here. In short, solid-state nuclear magnetic
resonance (NMR) spectroscopy has been used with the goal of picking up cis peptides in
proteins. NMR spectroscopy has a marked advantage over other techniques in that it
allows site-specific resolution. Consequently, NMR spectroscopy is a popular method for
studying structure and dynamics of proteins and other biological molecules.
6
Solution NMR has traditionally been the technique of choice for studying such
large molecules. In solution, rapid molecular tumbling averages out anisotropic
interactions, which results in sharp spectral lines, yielding good resolution. In solids,
crystallites are frozen in space, which causes anisotropic interactions to dominate, giving
broad spectra which are difficult to assign. However, with the advent of magic-angle
spinning (MAS) solid-state NMR, such problems have been virtually overcome, and it is
possible to obtain sharp, “solution-like” spectra in solid-state. With time, experimental
approaches have been developed which allow reintroduction of anisotropic couplings
during specified durations of a MAS solid-state NMR experiment. These interactions are
extremely useful for the measurement of distances and torsional angles.
In this thesis, a new solid-state NMR experiments shall be described which will
be sensitive to the conformation of a peptide about the N-C’ bond. In the next section, the
fundamental approaches of NMR spectroscopy shall be discussed. Chapter 2 shall discuss
the preparation of model compounds with trans- and cis-peptide bonds. The details of the
proposed experiment along with preliminary simulated results shall be described in
Chapter 3. Some preliminary experimental results on the model compounds are shown in
Chapter 4. Finally, future goals and possible modifications of the experiment will be
discussed in Chapter 5.
7
1.2
NMR Spectroscopy
1.2.1 Matter and spin
Matter is made up of atoms, which in turn are made up of electrons and nuclei. Each
atomic nucleus has a unique number of protons and neutrons. The number of protons
gives each nucleus their chemical identity, while the number of protons and neutrons
together determine the nucleus’s mass and magnetic properties. The latter arises out of a
special property, known as spin. Both proton and neutron has a spin of ½. The combined
effect of the spins of each of these particles determine the ground state nuclear spin value
for a given nucleus. It needs mention here that although nuclei can be excited to higher
energy states, conditions required to achieve the same are prohibitive, and we restrict our
discussion to nuclear ground states. Possible spin values for nuclear ground state are
either integral (0, 1, 2 …) or half integral (1/2, 3/2, 5/2, ….). In this thesis, we shall only
be considering nuclei whose nuclear spin has the value of ½.
The nuclei most typically used in NMR of peptides and proteins are 1H, 13C, and
15
N. While 1H is the most abundant isotope of hydrogen,
13
C and
15
N are extremely rare
in nature. Consequently, NMR signal cannot be detected for these nuclei in a natural
abundance sample. Samples have to be isotopically labeled in order for these nuclei to be
observed in an NMR experiment.
1.2.2 Zeeman splitting
A nucleus with spin I (as described above) is (2I + 1)-fold degenerate. When such a
nucleus is placed in a magnetic field, this degeneracy is lifted, and the (2I + 1) energy
8
levels are split. This is known as Zeeman splitting. For spin-1/2 (I = ½) nuclei, this
generates two energy levels, which we call α and β.
When a nucleus with non-zero nuclear spin I is placed in an external magnetic
field of strength B0, it interacts with the magnetic field with interaction energy (E) given
by:
(1)
where µ is the magnetic moment of the nucleus given by
(2)
where h is Planck’s constant (universal constant) and γ is the gyromagnetic ratio,
characteristic of the nucleus.
If the external field is assumed to be along the Z direction (in other words, if the direction
of the external field is defined as the Z-axis), then the Hamiltonian for the Zeeman
interaction can be written as:
(3)
where IZ is the spin angular momentum operator in the Z direction.
From this point, the external field B0 will be assumed to be along the Z-axis.
9
In other words, energy difference between the two states α and β is given by
equation (1). Thus, at a given temperature T and magnetic field strength B0, the relative
population of the two states α and β (denoted nα and nβ respectively) are related by
Boltzmann distribution as:
(4)
where kB is Boltzmann constant and E has been defined in equation (1).
For magnetic field strengths typically used in NMR experiments and temperatures
few fractions of a degrees K, the term inside parenthesis is extremely low, and a “hightemperature approximation” can be made wherein it can be written:
which can be rearranged to obtain the fractional population of the higher energy state
with respect to the ground state
(5)
At room temperatures and typical magnetic field strengths, this fraction is of the order of
10-5. This results in NMR being an insensitive technique. Nevertheless, this population
difference is sufficient to detect signals with reasonably good signal-to-noise ratio.
10
1.2.3 Fourier-Transform (FT) NMR
For spin-1/2 nuclei, equations (1) and (2) can be combined to write the difference in
energy between states α and β as:
(6)
where ω0 is known as the larmor frequency of the nucleus, and is fixed for a given
nucleus at a given magnetic field strength.
Short radiofrequency pulses generated by RF coils around the sample, applied at a
direction perpendicular to the external magnetic field (B0) will tilt the net magnetization
away from B0. This induces an oscillating current in the RF coil, which can be detected as
the signal. The signal is detected as a function of time (known as the free induction decay
or FID), and then fourier transformed to generate the NMR spectrum. In the most general
form, the FID can be written as:
(7)
where A is a constant depending on the experimental conditions, λ is damping constant,
and Ω0 = ω0 – ωref, where ωref is a reference larmor frequency. This signal can then be
fourier transformed as:
(8)
S(Ω) is the frequency-domain NMR spectrum.
11
It should be noted here that a pulse is described by a phase (Φ P), power (ωnut),
frequency (ωP) and duration (τP). The angle βP by which the magnetization is tipped away
from B0 given by:
(9)
where ωnut = |γBRF/2|, BRF being the oscillating RF field.
It should also be noted here that the RF pulse interacts with the nucleus most
strongly is the frequency ωP is exactly equal to the larmor frequency ω0. In this case, the
pulse is said to be “on resonance” with the nucleus. Each nucleus is said to “resonate at”
the larmor frequency.
1.2.4 Multidimensional NMR
Although the problem of insensitivity of NMR spectroscopy (see section 1.2.2) can be
overcome by using higher magnetic fields, the resolution is still not sufficient especially
for large molecules like polypeptides and proteins. Such molecules typically have a large
number of spins, and peaks from each of these make the spectrum extremely complicated
and difficult to assign. To overcome this problem, different spins can be correlated so as
to have additional dimensions in the spectrum. This was proposed by Jean Jeener 18 and
demonstrated
by Richard
Ernst
and
corworkers19.
Simplest
example
of
a
multidimensional NMR experiment is a 2D correlation experiment. In such experiments,
transverse magnetization is created on one nucleus (say 15N) and is frequency-labeled for
duration t1. This magnetization can then be transferred to another nucleus (say 13C). This
transfer is usually through J-coupling or dipolar coupling (the following section discusses
12
these interactions briefly). Following the transfer, the signal is collected (for time t2).
Thus, a two-dimensional FID is collected which can be double fourier transformed to
generate a 2D 15N-13C correlation spectrum:
In principle, higher dimensions are also achievable. Higher dimensions would increase
resolution to aid spectral assigning, while being expensive in terms of time.
Such multidimensional NMR experiment is extremely valuable for studying large
biomolecules, as it allows for site-specific resolution. This is a feature unique to NMR
spectroscopy, and gives it an advantage over other techniques commonly used for
structural studies of biomolecules.
1.2.5 Magnetic interactions
Having described the fundamentals of FT-NMR experiment, this section will describe the
various magnetic interactions that are typically present in a sample of spin-1/2 nuclei
placed in a magnetic field.
1.2.5.1 Chemical shift
From what was discussed in the previous section, it would appear that all nuclei of a
given type under a given magnetic field would have the exact same resonant frequency,
as larmor frequenct depends only on the nucleus (gyromagnetic ratio) and magnetic field
strength. However, this is not the case as the surrounding influences the resonant
frequency of the nuclei.
13
The external magnetic field B0 induces a current in the electronic clouds
surrounding the nucleus. This generates a microscopic magnetic field, Binduced, which can
be expressed as
(10)
where δ is the chemical shift tensor, a rank two tensor. It is to be noted here that
the symbol δ is used for the “deshielding” convention wherein resulting shift with respect
to reference larmor frequency is +δ. A similar “shielding” convention can be used, where
the symbol σ is used to describe a chemical shielding tensor, with opposite sign as the
chemical shift tensor. In the shielding convention, Binduced is written as:
In Cartesian coordinates, the chemical shift tensor can be written as:
For each nuclear site, it is possible to choose an axis frame such that the chemical shift
tensor becomes diagonal when expressed in the said axis frame. This axis frame is said to
be the principal axis frame of the said chemical shift tensor.
This interaction is small compared to the Zeeman interaction (see section 1.2.2),
and hence, it can be approximated to only have the δ ZZ term (note that in the principal
axis frame all off-diagonal terms are zero). Thus, the chemical shift Hamiltonian can be
written as:
14
(11)
This is known as the secular approximation. Here,
is the ZZ element of the
chemical shift tensor in the laboratory reference frame (the reference frame in which B0
field is along Z-axis), and θ defines the orientation of a particular crysyallite in a powder
sample, with respect to this frame.
Such orientation dependence means that all three principal components (δ XX, δYY,
and δZZ) of the chemical shift tensor can contribute to the chemical shift Hamiltonian,
depending on the oritentation of a given crystallite.
The chemical shift interaction can be separated into an isotropic and an
anisotropic component. As the name suggests, the isotropic component is independent of
the crystallite orientation. These components, denoted δ iso and δaniso can be written as:
The asymmetry parameter of the chemical shift, η, is given by:
(12)
In liquids, due to rapid molecular tumbling, the chemical shift is averaged to its isotropic
value. This is because when the molecular tumbling is fast on the NMR timescale, each
molecule can be assumed to have the same orientation with respect to B0.
However, in most solids, due to the absence of such tumbling, the chemical shift
has an orientational dependence. If the direction of B0 in the principal axis frame of the
chemical shift tensor is b0 and is described by the polar angles (φ, θ), then it can be
15
shown that the effective resonant frequency (ωCS) for a nucleus of larmor frequency ω0 is
given by:
which for axially symmetric chemical shift tensors (η = 0) can be written as:
(13)
As can be seen from equation (13), in a powder, each crystallite has a different
orientation with respect to B0, and consequently, experience slightly different chemical
shifts. Each crystallite has its peak at a slightly different location, which superimpose to
give a broad signal, known as “powder pattern”. This effect can be removed by magicangle spinning (see section 1.2.5). However, the oritentation-dependence of this
anisotropy also makes it a useful probe for measuring torsional angles. The potential of
chemical shift anisotropy (CSA) in measuring ω-angle in peptides and proteins will be
discussed Chapter 5.
1.2.5.2 Spin-spin interactions
A typical molecule contains more than one spins, each of which respond to the external
magnetic field (and experience chemical shift) as described in the previous section.
Additionally, each of these spins are coupled to each other. For spin-1/2 nuclei, the two
possible mechanisms of spin-spin coupling are J-coupling and dipolar coupling. For both
of these interactions, the Hamiltonian HC between nuclei i and j has the form:
16
(14)
where I is the spin angular momentum (given by I2 = Ix2 + Iy2 + Iz2) and subscripts i and j
denote the two nuclei, and C is a rank two tensor (similar to the chemical shift tensor
described in the previous section) describing the said interaction.
For each of these cases, random molecular tumbling in solution result in the
tensors being averaged to their isotropic value, which is defined by the trace of the tensor.
This is similar to the effect described for chemical shift, where molecular tumbling
reduces it to its isotropic value.
J-coupling
J-coupling (also known as scalar coupling or “through-bond” coupling) is the coupling
between two nuclei through bonding electrons. Their effect is strongest between two
nuclei which are directly bonded, and die away usually within 3 bonds. In solution NMR,
they cause splitting of NMR spectra. The J-coupling tensor has an anisotropic part which
is extremely small and can be neglected. The resulting secular-approximated Hamiltonian
is given by:
If the chemical shift difference between the two species is significantly larger than the
strength of the J-coupling, which is usually the case, then a further approximation (“weak
coupling” approximation) can be made and the Hamiltonian further simplified as:
While a common feature in solution NMR, they are usually dominated by stronger,
dipolar couplings (described in the next section), in solids. J-couplings are not used in
any of the experiments used in this thesis, and shall not be discussed any further.
17
Dipolar coupling
Dipolar coupling (also known as “direct” coupling or “through-space” coupling) is the
coupling between two nuclei without the involvement of electrons. The strength of the
coupling is directly related to the distance between the two nuclei (falls off as inversecubed of the distance). The dipolar tensor D is of the form:
(15)
where x, y and z are the projections of the internuclear vector on the coordinate axes, and
r is the length of the internuclear vector. Note that this is a traceless tensor, as the
diagonal elements add up to zero.
Using this in equation (13), it can be shown that the Hamiltonian for dipolar interaction
between nuclei i and j is given by:
(16)
where I is the spin angular momentum and subscripts i and j denote the two nuclei, e ij is
the unit vector parallel to the line joining the two nuclei, and bij is dipolar coupling
constant given by:
(17)
18
where µ0 is the permeability of free space, γi and γj the gyromagnetic ratios of the two
nuclei, and rij the distance between the nuclei.
Like chemical shift, the secular approximation can be used even for dipolar coupling to
simplify the interaction Hamiltonian (equation 13). This simplified Hamiltonian can be
written as:
is the secular dipolar coupling and θij is the angle
where
between the vector eij and the external magnetic field.
As for J-coupling, the weak coupling approximation can be made for dipolar
coupling to further simplify the Hamiltonian as:
(18)
As mentioned earlier, in liquids, rapid molecular tumbling averages this interaction to its
isotropic value, which is zero. Hence, dipolar coupling is usually not observed in
solution. However, in solids, due to restricted motion, the anisotropic part is not averaged
out. This is both a blessing and a nuisance. On one hand, this leads to “dipolar
broadening” of solid-state NMR spectra, a problem that can be removed by magic-angle
spinning (see section 1.5). On the other hand, dipolar coupling is a storehouse of
information. The dependence of the dipolar coupling constant on internuclear distance
(equation 16) makes dipolar coupling measurement a very useful tool for determining
structural restraints. Additionally, angular dependence of the secular dipolar coupling
constant makes it a useful probe for measuring torsional angles, a feature that has been
utilized in this work.
19
1.2.6 Solid-state NMR and Magic-angle spinning
The problems associated with anisotropic interactions, specifically, dipolar coupling and
chemical shift anisotropy, have been described above. These interactions result in the
spectral lines being too broad for meaningful assignment of the peaks. In order to obtain
high-resolution NMR spectra of solids, it is necessary to remove these effects. This is
achieved by magic-angle spinning.
As can be seen from equations (13) and (18) and keeping in mind the orientational
dependence of the term bij (equation 18), it can be seen that both of these terms have an
orientational dependence of the form (3cos2θ – 1). Here, θ is the angle between B0 and
orientation of the interaction tensor (chemical shift tensor and dipolar tensor
respectively). In a powder sample, θ takes all possible values as all molecular orientations
are possible.
If we spin this sample about an axis inclined at an angle θ R with respect to B0,
then θ varies with time as the molecule rotates. Under such circumstance, the average
orientational dependence, <3cos2θ -1> (angular brackets denoting averaged) can be
shown to be:
(19)
where β is the angle between the spinning axis and the Z axis of the principal axis frame
of the tensor.
θR, the angle between the sample and B0, is under the control of the experimenter.
As can be seen from equation (19), if θR is set to 54.74º, the term on the left hand side
20
becomes 0. This angle is called the “magic-angle”, introduced by Andrew et. al20. and
Lowe21. If the sample spinning is fast enough (so that θ is averaged rapidly compared to
the dipolar coupling and the chemical shift anisotropy), the anisotropic components are
averaged to zero, resulting in sharp spectral lines.
The terms “rotor period” is used to refer to the time taken by the sample to
complete one rotation.
1.2.7 Recoupling experiments
As mentioned above, anisotropic interactions like chemical shift anisotropy and dipolar
coupling, are both a blessing and a nuisance. While it is useful to remove their effect of
line broadening from solid-state NMR spectra, it would also be profitable to be able to
measure these quantities, as they contain useful structural information.
For this reason, techniques have been developed to reintroduce these interactions
under magic-angle spinning, for specific periods during the experiment. These are called
“recoupling” experiments. Carefully designed sequence of pulses can be used to
reintroduce chemical shift anisotropy, homonuclear dipolar coupling, and heteronuclear
dipolar coupling. Some of these techniques have been used in this thesis, and will be
described in Chapter 3. It is also possible to correlate two different anisotropic
interactions to measure the relative orientation of the two tensors, which is the essence of
measuring torsional angle using solid-state NMR. This is discussed in the next section.
21
1.2.8 Measuring torsional angles
Measurement of torsional angles is a key step to structural studies of proteins and
peptides. A polypeptide backbone has three different torsional angles, denoted φ, ψ and
ω. Two of these, φ and ψ, are needed to completely describe the three-dimensional
structure of the polypeptide chain. Consequently, more attention has been given to the
measurement of these angles. A number of magic-angle spinning solid-state NMR
experiments have been designed for measuring these torsional angles in proteins 22-31. This
section shall explain the way these experiments correlate two different anisotropic
interactions to give information about the relevant torsional angle.
Consider a molecular fragment consisting of
torsional angle about the
15
1
H-15N-13C-1H, in which the
N-13C bond is to be measured31. In this experiment,
15
N
transverse magnetization is generated, and dephased by reintroducing 1H-15N dipolar
coupling, in a time-resolved fashion. This dephased magnetization is frequency-labeled
(for site-specific resolution) and transferred to
13
C transverse magnetization. This
magnetization is then dephased by reintroducing 1H-13C dipolar coupling in a timeresolved manner, and finally the magnetization is detected. The result is a series of 2D
15
N-13C correlation spectra in which each cross-peak is modulated by the relative
orientation of the 1H-15N vector and the 1H-13C vector. The intensity of a given crosspeak is given by:
where DN is the dephasing of the
15
coupling, DC is the dephasing of the
N transverse magnetization due to 1H-15N dipolar
13
C transverse magnetization due to 1H-13C dipolar
coupling, and T(τmix) is the magnetization transfer efficiency for the 15N to 13C transfer.
22
Each of these functions depend on the relative orientation of the molecular
fragment with respect to B0. However, it is usually possible assume T(τmix) to be
constant, and consider only the two other functions. The functional forms of these are:
where ω is the orientation-dependent dipolar coupling being reintroduced ( 1H-15N amd
1
H-13C respectively). Hence, the intensity cross-peaks can be written as:
where ωHN and ωHC are the orientation-dependent dipolar couplings being reintroduced
for periods τN and τC respectively, and the angular brackets denote an average over all
crystallites.
The two dephasing periods τN and τC could be varied independently to generate
effectively a 4D experiment. However, time required for such an experiment would be
prohibitively long. Hence, the two dephasing periods are varied synchronously to
generate a 3D experiment.
This is the methodology that shall be used in this thesis for describing a new
experiment for obtaining information on geometry about the N-C’ bond in polypeptides.
In principle, different anisotropic interactions including dipolar couplings and chemical
shift anisotropy can be correlated for this purpose. These shall be discussed in Chapter 3,
and the most appropriate experiment shall be chosen from preliminary simulation results.
Chapter 4 shall discuss some of the results from the experiments performed on the model
compounds. Chapter 5 shall discuss some possible modifications to the experiments that
can be made for better results.
23
Chapter 2
Synthesis of model compounds
Linear diglycine (Figure 2.1) was chosen as a model compound with trans-peptide, and
2,5-diketopiperazine (Figure 2.2) was chosen as a model compound with cis-peptide.
Both of these compounds are commercially available. However, for our experiments, we
needed isotopically labeled (U-13C2,
15
N) versions of these compounds (see section
1.2.1). For this reason, both of these compounds were synthesized in-house. The
commercially available versions were purchased and used for standardization purposes.
This chapter describes the synthesis protocols used, optimazions required, and final
results obtained for the synthesis of these two compounds.
Figure 2.1: Glycylglycine
Figure 2.2: 2,5-diketopiperazine
All 1H NMR was taken at 400 MHz 1H larmor frequency and all 13C NMR was taken at
500 MHz 1H larmor frequency. All NMR spectra were recorded in D2O solvent unless
specified otherwise.
24
2.1 Synthesis of model compound with trans-peptide
Glycylglycine can be synthesized using solid-phase peptide synthesis using standard
Fmoc-chemistry. The original scheme that was planned for this synthesis is shown in
Figure 2.3.
Resin used was Wang Resin and was obtained from AAPPTec. Fmoc-
protected Glycine was also obtained from AAPPTec. DIC was obtained from Sigma
Aldrich and HOBt.H2O was obtained from AnaSpec Inc.
Figure 2.3: Proposed scheme for synthesis of glycylglycine
25
Glycylglycine was purchased from Bachem for standardization tests. 1H and 13C NMR of
this compound is shown (Figures 2.4 and 2.5 respectively).
Figure 2.4: 1H NMR of Glycylglycine (Bachem)
The following protocol was followed for the synthesis scheme shown in Figure 2.3. ~ 2 g
Wang Resin was washed with DCM for 30 minutes and 10% DMF in DCM for 30
minutes. 200 mg HOBt.H2O and 20 mg DMAP was added to Fmoc-protected Glycine
and dissolved in minimum amount of DMF. 200 µL DIC was added and allowed to stand
for 5 minutes. This mixture was then added to the resin and shaken for 14 hours. 0.1 mL
Pyridine and 120 µL Acetic anhydride was added to it and shaken for further 30 minutes.
26
The liquid was drained and the beads washed alternately with DCM and DMF (four times
each). After drying the beads were washed once more with DMF and 20% Piperidine in
DMF was added. It was shaken for 30 minutes. Kaiser test was performed after draining
the liquid and drying, to confirm the presence of free amine. The beads were again
washed alternately with DCM and DMF (four times each). After drying, it was washed
with 10% DMF in DCM.
Figure 2.5: 13C NMR of Glycylglycine (Bachem)
27
200 mg HOBt.H2O and 20 mg DMAP was added to Fmoc-protected Glycine and
dissolved in minimum amount of DMF. 200 µL DIC was added and the mixture was
allowed to stand for 5 minutes. This was added to the beads and shaken for 8 hours. The
liquid was washed alternately with DCM and DMF (four times each). 0.1 mL Pyridine
and 120 µL Acetic anhydride was added to it and shaken for further 30 minutes. The
liquid was drained and the beads washed alternately with DCM and DMF (four times
each). After drying the beads were washed once more with DMF and 20% Piperidine in
DMF was added. It was shaken for 30 minutes. Kaiser test was performed after draining
the liquid and drying, to confirm the presence of free amine. The beads were again
washed alternately with DCM and DMF (four times each). After drying, the beads were
washed once more with DCM 50% TFA in DCM was added and shaken for 2 hours. The
filtrate was collected. The beads were washed with small portions of TFA, and the was
combined with the filtrate.
DCM was evaporated at 35 ºC under vacuum, and the residue was dissolved in
appropriate solvent (usually D2O) for taking 1H and 13C NMR.
The product obtained was analyzed by taking 1H NMR (Fiugure 2.6) and
13
C NMR
(Figure 2.7). Clearly, Glycylglycine was not the major product. The product(s) present
could not be identified. Reason for not obtaining the product was not confirmed, but it
was probable that since the peptide was so short, there was possible cyclization (or other
side reactions) during cleavage from the resin.
28
Figure 2.6: 1H NMR of product obtained by proposed scheme
Figure 2.7: 13C NMR of product obtianed by proposed scheme
29
If cyclization was indeed the problem, then cleavage of the peptide from the resin
before the removal of the Fmoc group should solve the problem, as, in that case, there
would be no free amine group during cleavage which could cause partial cyclization.
What needed to be confirmed was whether cleavage could work in the presence of
Fmoc (most protocols have Fmoc groups removed before cleavage from resin). To check
this, Fmoc-protected Glycine was attached to the resin and then cleaved with 50% TFA
in DCM. 1H NMR of Fmoc-protected Glycine is shown in Figure 2.8 and that of the
product obtained is shown in Figure 2.9. These compounds are insoluble in D 2O, and
these NMR spectra were taken in CDCl3 solvent, with TMS added as internal reference.
Figure 2.9 is complicated as the compound is impure. However, presence of peak at 7.2
ppm confirms that the product contains Fmoc-group, which must have been cleaved from
the resin. In other words, it was confirmed that deprotection works in the presence of
Fmoc.
The next thing that had to be checked is, whether 50% TFA in DCM can, apart
from cleavage, also effect removal of the Fmoc group if treated for longer durations. This
would make the working up of the reaction much simpler, as DCM solvent is a lot easier
to remove than DMF, the solvent conventionally used for removal of Fmoc group.
To check this, the synthesis scheme (Figure 2.3) was repeated until the last two
steps. Then, the treatment with 50% TFA in DCM was continued for 4 hours, and the
product was isolated by evaportating DCM. 1H of this compound is shown in Figure 2.10.
30
Figure 2.8: 1H NMR of Fmoc-protected Glycine (AAPPTec) in CDCl3 with TMS standard
Figure 2.9: 1H NMR of Fmoc-protected Glycine after attachment to resin and cleavage, dissolved in
CDCl3 with TMS standard
31
Figure 2.10: 1H NMR of product obtained by treating resin-bound Fmoc-Gly-Gly with 50% TFA in
DCM, dissolved in CDCl3 with TMS standard
The 1H NMR does show a 1:1 peak corresponding to the two methylene groups.
However, comparison with 1H NMR of Glycylglycine (Figure 2.4) shows that the
position of these peaks are shifted by ~ 1 ppm. This was clearly not the desired product.
This clearly meant that this product had to be treated with piperidine in DMF for
the removal of the Fmoc group. However, the work-up of that step was a problem that
had to be solved. Normally, Fmoc removal is done while the peptide is bound to the resin
beads. After the reaction, solvent (containing excess reactant and unwanted product) can
be allowed to drain. However, if Fmoc removal was done after cleavage, purification of
the product would be difficult. HPLC purification could be done if the product could be
dissolved in water (Glycylglycine is soluble in water). However, that would require
removal of DMF first, which is quite difficult.
32
If it was possible to do Fmoc removal in DCM instead of DMF, purification
would be easier, as DCM could be easily removed. To check this, the synthesis was
repeated, with cleavage from resin done with Fmoc still attached. This compound was
treated with Piperidine in DCM. The 1H NMR (Figure 2.11) did not have the peaks
corresponding to the product (Figure 2.4). Clearly, Fmoc removal had to be done in
DMF.
ESI Mass Spectra was taken for this compound (Figure 2.12). The product could not be
identified even from its mass.
At this point, a method had to be found to remove DMF after reaction was over. Two
such methods were tried:
1.
Adding Toluene to create azeotrope:
Toluene was added to the solution in 1:5 ratio (i.e. 50 mL Toluene to be added to 10 ml
DMF). This creates an azeotrope. Evaportation at 60 degrees was found to dry the
solution in ~ 2 hrs. If it was still not dried, additional Toluene could be added and the
process could be continued to yield a dry mixture.
2.
Air-drying:
The solution was taken in a 2-necked or 3-necked round-bottomed flask. One of the
mouths was kept open and the rest (2 of them for 3-necked and 1 for 2-necked flasks)
33
were closed with septum. Through one of the septums (the only septum for 2-necked), air
was inject by inserting a syringe which is connected to a tube attached to the air vent. Air
flow was adjusted so that it is not high enough to spill the solution through the open
mouth. Dry mixture was obtained overnight or in two days depending on the volume of
the solvent.
Figure 2.11: 1H NMR of product obtained after cleavage in DCM
34
Figure 2.12: ESI Mass Spectra of product obtained after cleavage in DCM
Air-drying had the additional advantage of completely removing TFA. When Tolueneazeotrope was evaporated, the mixture still smelt of TFA. This is usually not reliable as
TFA can interfere with other reactions. After air-drying, there was no smell of TFA. For
this reason, this method of removing DMF was used henceforth.
The synthesis was repeated with the modifications described above. 1H and
13
C
NMR of the product is shown in Figures 2.13 and 2.14 respectively. As can be seen in the
1
H NMR (Figure 2.13), the 1:1 peaks are still shifted by ~ 1 ppm (compared to Figure
2.4). However, since it shows peaks corresponding to the Fmoc group, chances are that it
is Fmoc-Gly-Gly-OH.
To confirm this, ESI Mass Spectra was taken for this compound (Figure 2.15).
35
Figure 2.13: 1H NMR of product obtained with modified scheme
Figure 2.14: 13C NMR of product obtained with modified scheme
36
Figure 2.15: ESI Mass Spectra of product obtained with modified scheme
As can be seen, peak appears where it is expected for Fmoc-Gly-Gly-OH. This confirmed
that this is Fmoc-Gly-Gly-OH.
The only possible explanation for this is that, Fmoc removal becomes slower
when the peptide is bound to the resin. This makes intuitive sense as, when the resin is
bound, only the Fmoc group is oriented towards the outside, and it is easy for Piperidine
molecules to “find” the Fmoc groups. Whereas, when not bound to resin, the Fmoc-GlyGly-OH molecules are randomly oriented, and the Piperidine molecule has to “find” the
Fmoc group before the deprotection can take place.
37
With this contention, the process was repeated, this time the last deprotection
done for 4 hours and 24 hours. Overlaid 1H NMR of the products are shown (Figure 2.16)
As can be seen, 4 hour deprotection was still insufficient, while 24 hour deprotection
shows 1:1 methylene peaks where they are expected for glycylglycine (see Figure 2.4).
Also note that the product of 24-hour deprotection was insoluble in CDCl3 and hence was
dissolved in D2O for 1H NMR, which gives the strong D2O peak in the last product.
13
C NMR of this product (Figure 2.17) also matches well with that of
Glycylglycine (Figure 2.5). This method was used for the synthesis of (U-13C2,
15
N)
Glycylglycine. The scheme is shown in Figure 2.18.
The product, after air-drying, was dissolved in water and purified by RP-HPLC (Waters
1525) on a C18 column using water (with 0.01% TFA) and acetonitrile (with 0.01%
TFA) as solvent. The concentration of H2O was varied from 95% to 70% in 20 minutes
(8mL/min flow rate). Absorbance at 200 nm is shown (Figure 2.19). The peak at 10
minutes is confirmed to be Glycylglycine by 1H NMR obtained by lyophilizing the
fraction and dissolving in D2O. Overlaid spectra of this compound with that of
commercially available Glycylglycine (Bachem) is shown (Figure 2.20).
38
Figure 2.16: Overlaid 1H NMR of product obtained with modified scheme, with Fmoc removal done
for 30 mins, 4 hours and 24hours.
39
Figure 2.17:
13
C NMR of product obtained with modified scheme, with Fmoc removal done for 24
hours
40
Figure 2.18: The optimized synthesis scheme that was followed for the synthesis of (U-13C2,
Glycylglycine
41
15
N)
Figure 2.19: RP-HPLC of Glycylglycine at 220 nm
Figure 2.20: Overlaid 1H NMR of
Glycylglycine (Bachem) and Glycylglycine prepared by the
optimized scheme
42
The product was crystallized as its HCl salt. Needle-like crystals appeared in 2 hours
(Figure 2.21).
Figure 2.21: HCl crystal of Glycylglycine
2.1.1 Problems associated with solid phase peptide synthesis
While being being reasonably simple, solid-phase peptide synthesis had one problem for
our purposes. Usually, for such synthesis, the protected amino acids are not the most
expensive reagent. Hence, most protocols can afford to follow a simple trick: in every
step, 3-5 folds of excess amino acid is added, with respect to the resin loading capacity.
This is to ensure that all the sites on the resin (or the peptide already attached) are
43
occupied by the amino acid, so as to maximize the yield. For our purpose, the isotopically
labeled amino acid was indeed the most expensive reagent, and it was not possible to add
too much excess. This meant that after the first addition, there would probably be a
significant number of sites on the resin that would be unoccupied. Although Pyrdine and
Acectic anyhydride was added to cap the unoccupied sites, it still meant that during the
next attachment, some of the Fmoc-protected glycine would attach directly to the resin
(unbound sites) rather than to the first Glycine. This, apart from reducing the yield, would
cause the resultant compound to be impure, as there will be some Glycine contamination.
This was a problem that could not be solved for the synthesis of (U-13C2,
15
N)
Glycylglycine.
2.1.2 Synthesis of (U-13C2, 15N) Glycylglycine
(U-13C2, 15N) Fmoc-Glycine was purchased from Cambridge Isotopes Ltd. 2.0021
g Wang Resin (AAPPTec) was washed with DCM for 30 mins and 10% DMF in DCM
for 38 mins. 200.6 mg HOBt.H2O and 20.7 mg DMAP was added to (U-13C2, 15N) FmocGly-OH (Cambridge Isotopes Ltd.) and dissolved in minimum DMF. 200 μL DIC was
added and allowed to stand for 5 mins. This mixture was added to the resin and shaken
for 14 hrs. 0.1 mL pyridine and 120 μL Ac2O was added and it was shaken for further 30
mins. The liquid was drained and the beads were washed alternately with DCM and DMF
four times. After drying, beads were washed once more with DMF, and 20% piperidine
in DMF was added. It was shaken for 30 mins. Kaiser test was done after drying, and it
showed presence of free amine. The beads were again washed alternately with DCM and
DMF four times. After drying, it was ashed with 10% DMF in DCM.
44
212.4 mg HOBt.H2O and 13.9 g DMAP was added to (U-13C2,
15
N) Fmoc-Gly-
OH (Cambridge Isotopes Ltd.) and dissolved in minimum DMF. 200 μL DIC was added
and allowed to stand for 5 mins. This was added to the beads and shaken for 8 hrs. The
liquid was drained and washed alternately with DCM and DMF four times. After drying,
50% TFA in DCM was added and shaken for 2 hrs. The filtrate was collected. The beads
were washed with small volumes of TFA and the filtrate was combined.
TFA was removed by evaporating under vacuum first at 60 ºC for 2 hrs, then at 35
ºC overnight. For complete removal of TFA, the mixture was air-dried for 24 hrs.
20 % piperidine in DMF was added to the residue and shaken for 24 hrs. The
reaction mixture was air-dried, and the residue was dissolved in water and purified by
RP-HPLC (Waters) with H2O-Acetonitrile solvent, with H2O concentration varying from
95% to 70% in 20 mins (8mL/min).
The purified product was lyophilized and crystallized as its HCl salt. 1H NMR in
D2O (Figure 2.22), apart from splitting due to labellings introduced, showed traces of
impurity which is probably Glycine, introduced for reasons explained in the previous
section. This might also be the reason why the crystal quality (Figure 2.23) was inferior
compared to natural abundance Glycylglycine (Figure 2.21).
45
Figure 2.22: 1H NMR of (U-13C2, 15N) Glycylglycine
Figure 2.23: HCl crystal of (U-13C2, 15N) Glycylglycine
46
2.2
Synthesis of model compound with cis-peptide
2.2.1 Synthesis of 2,5-diketopiperazine using solid phase peptide synthesis
Cylcic peptides can synthesized using solid-phase peptide synthesis. On-resin cylcization,
which involves cyclization of the peptide while being bound to the resin, has been
reported in literature32,
33
. Taylor and coworkers34 have prepared three simple
piperazinediones using Kaiser oxime resin, and Smith et. al35. have described the protocol
for synthesizing cyclic dipeptides using on-resin cyclization on Kaiser oxime resin.
However, Kaiser oxime resin is not compatible with Fmoc chemistry, and Boc
chemistry has to be used. Boc chemistry has the following disadvantages:
1.
Reactions are extremely slow. The attachment of the first peptide to the resin
takes close to 24 hours (12 hours for Fmoc chemistry on Wang resin), and the subsequent
peptides require 12 hours (4-5 hours for Fmoc chemistry).
2.
Removal of Boc group is extremely difficult. It requires 95% TFA which is
extremely dangerous. It is also a very slow reaction, requiring ~ 16 hours (Fmoc group
can be removed in 2 hours using 50% TFA).
Additionally, solid-phase peptide synthesis has the problems described in the previous
section. For these reasons, 2,5-diketopiperazine was not synthesized using solid-phase
peptide synthesis. Instead, a non-conventional approach was used, which is described in
the following section. This method is extremely simple, very fast (2,5-diketopiperazine
can be obtained from glycine in less than an hour), and the product extremely pure.
47
2.2.2 Synthesis of 2,5-diketopiperazine using microwave-asisted synthesis
Microwave-assisted synthesis of 2,5-diketopiperazine has been described36. Briefly, 250
mg Glycine was taken in a 10 mL microwave reaction vessel, and 2 mL DMF was added.
It was subjected to microwave reaction in CEM Discover SP microwave reactor. Set
point temperature was set at 210 ºC and the hold time was varied to get maximum yield.
The product can be distinguished from unreacted Glycine by
13
C NMR taken in
D2O. While carbonyl of Glycine shows peak at ~172 ppm, 2,5-diketopiperazine shows
peak at ~ 168 ppm. When the hold time is increased, the peak at 168 ppm is seen to
increase in intensity at the cost of the peak at 172 ppm.
Initial reactions were done with hold times of 3, 6, and 9 mins (Figure 2.24). Even
at 9 mins, considerable amount of Glycine was uncreacted. Next, reactions were done
with hold times of 15, 30 and 45 mins (Figure 2.25). It was seen that at 30 mins, all the
glycine converted to 2,5-diketopiperazine.
Comparison of this spectrum with that for control 2,5-diketopiperazine (obtained
from Sigma Aldrich) confirmed the formation of the product (Figure 2.26). 1H NMR of
the same compound is shown (Figure 2.27). Reaction profile for this reaction is shown
(Figure 2.28).
48
Figure 2.24: Overlaid
13
C NMR of product obtained by microwave-assisted synthesis of 2,5-
diketopiperazine, with hold times of 3, 6 and 9 minutes
Figure 2.25: Overlaid 13C NMR spectra of product obtained by microwave-assisted synthesis of 2,5diketopiperazine with hold times of 15, 30 and 45 minutes
49
Figure 2.26: Overlaid 13C NMR of 2,5-diketopiperazine (Sigma) and 2,5-diketopiperazine synthesized
by microwave-assisted synthesis
Figure 2.27: 1H NMR of 2,5-diketopiperazine synthesized by microwave-assisted synthesis
50
Figure 2.28: Reaction profile for microwave-assisted synthesis of 2,5-diketopiperazine
Although the NMR spectra looked clean, the product didn’t crystallize. So, it was
passed through HPLC column under conditions similar to purification of Glycylglycine.
As a control experiment, 2,5-diketopiperazine purchased from Sigma Aldrich was also
passed through HPLC column in a similar fashion. Overlaid HPLC at 220 nm for control
2,5-diketopiperazine (obtained from Sigma Aldrich) and 2,5-diketopiperazine prepared
by microwave-assisted synthesis is shown in Figure 2.29. This confirmed the product
eluting at 10 mins was 2,5-diketopiperazine. It was further compared by 13C NMR of the
product after lyophilizing the fractions (Figure 2.30).
51
Figure 2.29: Overlaid HPLC at 220 nm of 2,5-diketopiperazine (Sigma) and 2,5-diketopiperazine
synthesized by microwave-assisted synthesis
Figure 2.30: Overlaid 1H NMR of 2,5-diketopiperazine (Sigma) and 2,5-diketopiperazine synthesized
by microwave-assisted synthesis after HPLC purification
52
2.2.3 Synthesis of (U-13C2. 15N) 2,5-diketopiperazine
(U-13C2,15N) Glycine (purchased from Cambridge Isotopes Ltd.). 250 mg (U-13C2,15N)
Glycine was taken in a 10 mL microwave reaction vessel, and 2 mL DMF was added. It
was subjected to microwave reaction in CEM Discover microwave reactor. Set point
temperature was set to 210 ºC. The hold time was kept at 30 mins. The reaction profile
for this reaction is shown (Figure 2.31).
The product was purified by HPLC as described above. The purified product
showed splitting in 13C NMR (Figure 2.32) as expected due to the labeling introduced.
Figure 2.31: Reaction profile for the microwave-assisted synthesis of (U-13C2,
diketopiperazine
53
15
N) 2,5-
Figure 2.32: 13C NMR of (U-13C2, 15N) 2,5-diketopiperazine
54
Chapter 3
Design of NMR experiment
Section 1.2.8 described the essence of torsional angle measurement using solid-state
NMR spectroscopy. Such experiments extensively use dipolar recoupling experiments
(see section 1.2.7). The proposed experiments, too, shall make use of such recoupling
techniques, and a more detailed discussion of these techniques shall be presented here.
3.1
3.1.1
Recoupling experiments
Heteronuclear dipolar recoupling
Techniques for measuring heteronuclear dipolar recoupling have a similar form. All of
them are based on a spin echo experiment, which normally refocuses heteronuclear
dipolar couplings. However, in these techniques, specially designed sequence of rf pulses
are applied on one of the spins to prevent complete refocusing of the dipolar coupling. In
this thesis, two different dipolar recoupling techniques shall be discussed: 13C-15N dipolar
recoupling and 15N-1H dipolar recoupling.
13
C-15N dipolar recoupling
The most common and robust technique for
13
C-15N dipolar recoupling is
Rotational Echo Double Resonance (REDOR)37. The pulse sequence for this technique is
shown in Figure 3.1.
REDOR consists of two trains of pulses applied on the
15
N channel, which are
rotor-synchronized. Pulses are applied every half rotor period on the
55
15
N channel. The
two trains of pulses are separated by a period of two rotor periods, which contain, at the
center, the echo pulse on the
13
C channel. The
13
C-15N dipolar coupling evolves only
during the train of 15N pulses, known as the REDOR “mixing period”, denoted by t mix.
The effect of the
13
C-15N dipolar coupling is to dephase the
13
C transverse
magnetization. This in turn reduces the 13C peak intensity of the resultant NMR spectrum.
Figure 3.1: Pulse sequence for Rotational Echo Double Reosnance (REDOR) experiment 37
In practice, a series of experiments are performed where t mix is incremented in
steps of 2τR where τR is the rotor period. The
13
C peak intensity as a function of the
REDOR mixing period is then plotted to give a REDOR dephasing pattern. Experimental
REDOR dephasing patterns can be fitted to simulated patterns to extract the the
13
C-15N
internuclear distance. A simulated REDOR experiment with 13C-15N distance of 1.5 Å is
shown in Figure 3.2
56
Figure 3.2: Simulated 13C-15N REDOR dephasing. Simulation done using SPINEVOLUTION38
15
N-1H dipolar recoupling
Most
solid-state
NMR
spectroscopy
experiments
biomolecules
involve
of
1
H
decoupling following initial 1H
flip and CP transfer. This is
because the large number of 1H’s
typically present in a biomolecule
are involved in strong dipolar
Figure 3.3: 15N-1H dephasing pattern. Figure taken from
reference 39
coupling network, which are not
removed completely even after
magic-angle spinning. Such decoupling techniques, apart from removing homonuclear
57
1
H-1H couplings, also remove heternuclear couplings of 1H with neighboring
15
13
C and/or
N.
An experiment like REDOR cannot be performed on
15
N-1H coupled systems for
this reason, unless 1H-diluted proteins are prepared, which is beyond the scope of this
thesis. The method employed for recoupling
15
N-1H dipolar interaction has been
described39 in literature. It uses a special decoupling on 1H for specific duration of the
experiment. Such special decoupling only removes homonuclear
1
H-1H dipolar
interaction while allowing heteronuclear dipolar interactions to evolve for the specified
duration.
15
N-1H dipolar coupling is stronger than
13
C-15N dipolar coupling. This can be
easily concluded by looking at the expression for dipolar coupling constant (Equation 17)
and noting that gyromagnetic ratio of 1H is much higher than that of 13C. For this reason,
15
N-1H dephasing curve decays much faster than 13C-15N decay, as shown by Figure 3.3,
which is taken from reference 39. As can be seen,
0.25 ms, whereas
13
15
N-1H dephasing decays to zero in ~
C-15N dephasing decays to zero in ~ 2 ms (Figure 3.2). This feature
has been utilized and is discussed in Chapter 5.
3.1.2 Homonuclear dipolar recoupling
For the purposes of this thesis, discussion on homonuclear dipolar recoupling shall be
restricted to recoupling of 13C-13C dipolar interaction. Unlike 13C-15N dipolar interaction,
there is no one “best” technique for reintroducing
13
C-13C dipolar interaction. Since the
first of its kind40 known as Dipolar Recovery At the Magic Angle (DRAMA), a series of
58
such methods have been published41-45 which are either entirely new or modification of a
previously published method.
From these choices, four methods were chosen as probably candidates for the
purpose of the current work. Dipolar Recovery with A Windowless Sequence
(DRAWS42),
MELding
Of
spin-locking
and
DRAMA
(MELODRAMA41),
Permutationally Offset STabilized C7 (POST-C744) and Supercycled POST-C5 (SPC-545)
experiments were chosen for their higher bandwidth and/or lower rf power requirements.
The best candidate, to be ultimately used in the experiment, was chosen based on the
sensitivity of the experiment, as obtained from simulations, described in section 3.2.
The rf power requirement plays an important role. All of these techniques require
the rf power on the 13C channel, in frequency units, to be a certain multiple of the sample
spinning frequency. However, the rf power on the 1H channel for decoupling should be ~
50 kHz higher than the
13
C channel power for effective decoupling. Taken together, a
higher power requirement for the
13
C-13C recoupling technique would indirectly affect
the power requirements on the 1H channel. Too high rf powers generates heat and are
capable of damaging the sample and the NMR probe.
Another point to be kept in mind is that these techniques also vary in terms of
how well they compensate for chemical shift anisotropy. This is important as carbonyl
carbon, a part of the spin system we are interested in, usually has high chemical shift
anisotropy.
59
3.1.3 Recoupling of chemical shift anisotropy
Like dipolar coupling, chemical shift anisotropy is also averaged to zero by magic angle
sample spinning. However, this too can be reintroduced by specially designed pulse
sequence46 .
While high chemical shift anisotropy of carbonyl could be a nuisance when
reintroducing
13
C-13C dipolar interaction, it itself could provide an alternative way of
obtaining information about the geometry of the peptide bond. A change in the ω-angle
would cause a change in the relative orientation of the
13
C-15N internuclear vector with
respect to the chemical shift tensor of the carbonyl carbon. Thus, measuring the relative
orientation of these two anisotropic interactions could also prove fruitful in measuring ωangle. The possibility of using this method is discussed in Chapter 5.
3.2
Cα-N-C-Cα experiment
The ω-torsinal angle between residues n and (n+1) is defined by the nuclei αCn+1, Nn+1,
C’n, and αCn. This angle can be thought of as the angle between the αCn+1- Nn+1 bond and
the OCn- αCn bond, which can be measured by the correlated evolution of αCn+1- Nn+1 and
C’n- αCn dipolar coupling. A schematic representation of the pulse sequence which can
achieve this goal is shown in Figure 3.4.
60
Figure 3.4: Schematic representation of the pulse sequence for measuring relative orientation of αC-N
and C’-αC internuclear vector
Briefly, after initial 1H flip and CP transfer to 13C to create transverse 13C magnetization,
13
C-15N dipolar coupling is reintroduced by REDOR pulse sequence (see section 3.1.1).
Following this, the magnetization is transferred to
15
N and then back to 13C. The last CP
transfer is selective, so that only the carbonyl carbons are magnetized. This is ensured by
adjusting the frequency offset of the CP spin-locking pulse and proper phase cycling. At
this point,
13
C-13C dipolar coupling is reintroduced by an appropriate pulse sequence.
This results in 13C magnetization which has been dephased by the correlated evolution of
13
C-15N and
13
C-13C magnetization. This magnetization is then detected as FID (see
section 1.2.3) and fourier transformed to generate an 1D
13
C spectrum. A series of such
spectra are generated where the duration of the recoupling periods are increased
synchronously, as described in section 1.2.8. The intensity of the
61
13
C peak as a function
of the recoupling period can be fitted to simulation to extract the relative oritentation of
the said interactions.
If this experiment is to be performed on a polypeptide, an additional dimension is
necessary for spectral assignment. This can be achieved by introducing a chemical shift
evolution period on
15
N (Figure 3.5). This frequency-labeled magnetization can then be
transferred to carbonyl before
13
C-13C recoupling and detection. This signal after double
fourier transform will result in a 2D
13
C-15N correlation spectra. A series of such spectra
with synchronously varying recoupling periods can be collected, where the intensity
variation of each peak can be fitted to simulations.
Figure 3.5: Schematic representation for same experiment with increased dimensionality
3.2.1 Choice of C-C recoupling technique
As mentioned in section 3.1.2, out of four homonuclear dipolar recoupling experiments,
DRAWS, MELODRAMA, POST-C7 and SPC-5, the one best suited for this experiment
was to be chosen. For this, each of these experiments was simulated using
62
SPINEVOLUTION34. Each simulation was performed on both trans and cis geometry.
The geometry was defined by Cartesian coordinates of the four nuclei ( αCn+1, Nn+1, Cn’,
and
α
Cn), obtained by generating the glycylglycine with trans and cis geometry,
respectively, in PyMol47. The simulations were performed with magnetic field strength
500 MHz 1H larmor frequency and 11.904 kHz magic angle spinning (so that the rotor
period was 84 µs).
The results from the simulations are shown in Figure 3.6, (a)-(d) represent,
respectively, DRAWS, MELODRAMA, POST-C7 and SPC-5 correlated evolution with
REDOR. As can be seen, DRAWS (Figure 3.6a) yields little difference in the dephasing
between trans and cis geometry. MELODRAMA (Figure 3.6b) yields some visible
change, and the key feature of the expected difference in the trajectory is already present
here. The trajectory of the trans geometry, after the initial decay, becomes flat during the
region of ~ 1 ms – 1.5 ms of evolution time, whereas the same for cis decays steadily till
~ 3 ms.
This feature is more dominant with POST-C7 and SPC-5 (Figures 3.6c and 3.6d
respectively). Here, from the region of ~ 0.5 ms – 1 ms of evolution time, the trans
trajectory flattens out while the cis trajectory shows a minima. While this is only a
preliminary result on which more work needs to be done (described in section 3.2.2), this
shows the direction in which to proceed.
Although both POST-C7 and SPC-5 yielded trajectories that looked similar,
POST-C7 was chosen as it had a higher recoupling efficiency compared to SPC-5.
63
3.2.2 Choice of ratio of mixing times
The next factor that had to be fixed was the ratio of mixing times that was to be used. As
was described in section 1.2.8, experiments for measuring torsional angles, in the most
general form, are 4-dimensional experiments (three dimensional if we omit the chemical
shift evolution dimension, as in this case). This would be the case if the two anistropic
evolutions were allowed to evolve independently. However, to cut down on the time
requirements, we vary the two dephasing periods synchronously. This means that the two
mixing times can only be varied at a fixed ratio. This immediately raises a question as to
what should this ratio be.
Figure 3.6: Simulations for the choice of 13C-13C recoupling element. (a) DRAWS (b) MELODRAMA
(c) SPC-5 (d) POST-C7
64
Such correlation experiments are most sensitive when maximum interference
between the two evolutions can be achieved. This happens if both the interactions
dephase to their respective minima at the same time. Figure 3.7 shows the simulated
dephasing curve of both REDOR and POST-C7 performed on the same molecular
fragment under similar conditions (αC-N-C’-αC fragment of trans glycylglycine, 500
MHz 1H larmor frequency, and 11.904 kHz magic angle spinning).
Figure
3.7:
Simulated
dephasing of
REDOR and
SPINEVOLUTION38
65
POST-C7.
Simulation
done
using
As can be seen, POST-C7 decays to its minima much faster than REDOR. This is
expected, as
13
C-13C dipolar coupling is much stronger compared to
13
C-15N dipolar
coupling (which, once again, can be related to the gyromagnetic ratios of the two nuclei).
Intuitively, it appears that allowing REDOR to evolve for twice as long as POSTC7 would result in the maximum interference. Indeed, in the plots shown in Figure 3.6,
REDOR mixing period was twice as much as POST-C7 mixing period. To verify if this
really is the most sensitive ratio, two more simulations were performed, one where
REDOR mixing period was kept equal to POST-C7 mixing period, and another where
REDOR mixing period was kept half of POST-C7 mixing. The simulated dephasing
patterns are shown in Figure 3.8. It is to be noted that the X-axis has been labeled by
POST-C7 mixing time in all the three plots shown.
As can be seen, the most sensitive experiment is the one in which REDOR mixing
period is kept twice that of POST-C7 mixing period. However, this is a parameter that is
not too difficult to vary while running the actual experiments, and might be worthwhile to
actually verify these predictions experimentally.
At this point, we wanted to verify if we could, in addition to distinguishing cis and trans
peptide bonds, also measure deviations from planarity of a peptide. To this end, similar
experiment was simulated (with REODR mixing period kept at twice POST-C7 mixing
period) on different geometries. Specifically, the αC-N-C’-αC torsional angle was varied
from 180º (completely planar) to 150º (30º deviation from planarity), in steps of 10º. The
simulated curves are shown in Figure 3.9
66
Figure 3.8: Simulation for the choice of ratio of REDOR and POST-C7 mixing time. (a) REDOR
mixing time = POST-C7 mixing time (b) REDOR mixing time = 2 * POST-C7 mixing time (c)
REDOR mixing time = 0.5 * POST-C7 mixing time
67
Figure 3.9: Sensitivity of simulated dephasing to variation of ω-torsional angle
As can be seen from the simulations, sensitivity of the experiment to small deviations of
the ω-torsinal angle is insignificant. This means, with the current experiment, it would be
unreasonable to hope to measure deviations from planarity of the peptide bond. However,
other modifications of this experiment are possible, and are discussed in the following
section, which might improve sensitivity.
3.3
Utilizing carbonyl CSA for ω-angle measurement
As discussed in section 1.2.5.1, chemical shift has an anisotropic component, called
chemical shift anisotropy (CSA). In principle, this interaction should also be exploitable
for torsional angle measurements. We shall focus on CSA of the carbonyl carbon as its
68
magnitude is much higher than any of the other nuclei relevant here. For our work, it can
be seen that the relative orientation of the carbonyl CSA with respect to the αC-N bond is
changed when the ω-torsinal angle changes. Hence, it should be possible, in theory, to
correlate αC-N dipolar coupling evolution with carbonyl CSA to obtain information about
the peptide bond geometry.
Schematic representation of the pulse sequence for correlating αC-N dipolar
coupling with carbonyl CSA is shown in Figure 3.10
Figure 3.10: Schematic representation of the pulse sequence for measuring the relative orientation of
the αC-N bond with carbonyl CSA
69
This is similar to the pulse sequence shown in Figure 3.4 with the C-C recoupling period
being replaced by the CSA recoupling period. Note that the magnetization prior to this
second recoupling period has been transferred selectively to the carbonyl carbons.
Chemical shift anisotropy is removed under the effect of magic-angle spinning. It
can be reintroduced by effect of rf pulses, as described by Chan and Tycko 46. It has to be
kept in mind that when simulating CSA, the CSA parameters, i.e. the anisotropy (δ aniso)
and asymmetry (η) must be known a priori, along with the set of (α, β, γ) angles that
define the orientation of the CSA with respect to the laboratory frame. For the present
work, these values were taken from the work of Karlsson et. al48. They reported CSA
parameters for six compounds. The compound they label as compound V, a tripeptide
AGG, is similar to glycylglycine, the model compound that has been used in this work.
They have reported the CSA parameters for the carbonyl of the Glycine at the center
(denoted as ‘k’ in their paper).
Using these values, the ROCSA dephasing was simulated in a similar manner.
The simulated curve is shown in Figure 3.11.
The feature that attracts attention here is the timescale of the decay. As can be seen, the
magnetization decays in ~ 0.25 ms. This is almost an order of magnitude shorter than the
timescale of decay of the REDOR experiment, ~ 1.5 ms (see Figure 3.2 and Figure 3.7).
70
Figure 3.11: Simulated decay curve for the ROCSA experiment46
As mentioned earlier, the experiment performs the best if the decay times of the two
sequences are very similar. While in principle this technique could still be made to work
by keeping the REDOR mixing time 10 times of the ROCSA mixing time, it should also
be kept in mind that after 10 REDOR cycles most of the magnetization will decay due to
relaxation processes which have not been considered in the simulations.
We speculate that this problem would be circumvented if 15N-1H dipolar coupling
was used instead of 13C-15N dipolar coupling. Relative orientation of 15N-1H internuclear
vector with carbonyl CSA is affected in the same manner as the relative orientation of the
C-N internuclear vector with carbonyl CSA. However, the additional advantage here is
that
15
N-1H dipolar recoupling decays in the same timescale as carbonyl CSA, as can be
71
seen from Figure 3.3 (Figure 2 of reference 39). A schematic representation of the
proposed experiment is shown in Figure 3.12.
Figure 3.12: Schematic representation of the pulse sequence for measuring the relative orientation of
15
N-1H internuclear vector and carbonyl CSA
72
Chapter 4
Preliminary results
This section shall describe some of the preliminary results obtained from solid-state
NMR experiments on the model compounds. While the complete picture is has still
proved elusive, results from some control experiment are worthwhile and give a clear
direction for progress. All experiments described here were performed under magnetic
field strength of 500 MHz 1H larmor frequency. Magic angle sample spinning rate was
11.904 kHz (rotor time of 84 µs).
4.1
Solid-state 1D CP-MAS experiments
The isotopically labeled model compounds with trans- and cis-peptide bonds were
synthesized, purified, and crystallized, as described in Chapter 2. Both of these crystals
were packed into Varian T3 HXY 3.2 mm MAS probe by centrifugation. As a first step,
1D
13
C and
15
N solid-state CP-MAS spectra was obtained for both the samples. These
spectra are shown in Figures 4.1-4.4.
As can be seen, 1D
15
N spectra of Glycylglycine (Figure 4.2) shows an extra
peak, probably arising from some impurity (which is presumably glycine, as explained in
section 2.1.1). However, this does not pose a significant hindrance for the purposes of
this experiment. This is because, all the pulse sequences proposed for this experiment
involves CP transfers from 13Cα to 15N and then from 15N to 13C’. This is possible only in
molecules containing a αC-N-C’ bond (peptide bond). Thus, only signal that would be
seen in this experiment would arise out of compound(s) containing at least one peptide
bond. Hence, small Glycine contamination is insignificant.
73
Also to be noted are the spinning sidebands of the carbonyl peak in 1D
13
C CP-
MAS spectrum, for both Glycylglyine (Figure 4.1) and 2,5-diketopiperazine (Figure 4.2).
This is due to high chemical shift anisotropy of carbonyl carbons, which is not
completely removed at 11.904 magic-angle spinning. Spinning sidebands can be easily
recognized as they are separated from the central peak (in frequency units), on either side,
by an integral multiple of the spinning frequency. The spinning sidebands seen here are
separated by 11.904 kHz (n=1 sideband).
Although higher spinning speeds can eliminate such spinning sidebands, our
experimental requirements do not allow us to spin our sample too high, as discussed in
section 3.1.2. However, these sidebands are easy to detect and low in intensity, and hence
does not cause too much of a hindrance.
Figure 4.1: 1D 13C CP-MAS spectrum of (U-13C2,
15
MHz and 11.904 kHz magic-angle-spining
74
N) Glycylglycine at magnetic field strength 500
Figure 4.2: 1D 15N CP-MAS spectrum of (U-13C2,
15
N) Glycylglycine at magnetic field strength 500
MHz and 11.904 kHz magic-angle-spinning
Figure 4.3: 1D 13C CP-MAS spectrum of (U-13C2, 15N) 2,5-diketopiperazine at magnetic field strength
500 MHz and 11.904 kHz magic-angle-spinning
75
Figure 4.4: 1D 15N CP-MAS spectrum of (U-13C2, 15N) 2,5-diketopiperazine at magnetic field strength
500 MHz and 11.904 kHz magic-angle-spinning
The next step to proceed with the proposed experiment was to optimize the CP conditions
for 13C to 15N and then from 15N selectively to 13C (carbonyl). The frequency offset of the
spin-lock pulse for the selective CP and the phase cycling had to be set such that only
carbonyl peak was seen in the 1D 13C spectrum. This was obtained for Glycylglycine, as
shown in Figure 4.5. Note that the singal-to-noise is poorer (compared to Figure 4.1), as
is expected, as there is some signal loss during CP transfer.
Similar 1D 13C spectrum was obtained for 2,5-diketopiperazine (not shown).
76
Figure 4.5: 1D
13
C spectrum of Glycylglycine, after CP transfers to
15
N and selectively to
13
C
(carbonyl)
Intensity of this carbonyl peak can now be followed as a function of REDOR mixing time
or POST-C7 mixing time. Once both the experiments have been calibrated, the two
elements (REDOR and POST-C7 respectively) can be put together to run the complete
sequence.
4.2
REDOR experiments
The REDOR experiment was performed on both the model compounds, along with the
two cross-polarization transfers, the conditions for which had already been optimized.
The intensity of the carbonyl peak (Figure 4.5) was followed as a function of REDOR
77
mixing time. This dephasing pattern, along with simulated REDOR curve is shown in
Figure 4.6 (Glycylglycine) and Figure 4.7 (2,5-diketopiperazine). For the simulation
(performed using SPINEVOLUTION38,
13
C-15N distance was set to 1.35 Å for both the
molecules for best fitting.
Note that in these simulations, relaxation has not been included. Yet, the fitting
with experimental data is quite good, as can be seen from the plots (Figures 4.6 and 4.7).
Figure 4.6: Experimental (red dot) and simulated (blue line) REDOR dephasing pattern of
Glycylglycine. Simulation done with 13C-15N distance set to 1.35 Å
78
Figure 4.7: Experimental (red dot) and simulated (blue line) REDOR dephasing pattern of 2,5diketopiperazine. Simulation done with 13C-15N distance set to 1.35 Å
79
Chapter 5
Conclusions and future work
Accurate determination of protein structure is an important aspect of describing the
biological function of a protein. The three dimensional structure of a protein backbone is
completely described by the various torsional angles. This can be realized by considering
the protein backbone as a chain of N atoms, which has 3N-6 degrees of freedom. These
are N-1 bond lengths, N-2 bond angles, and N-3 torsional angles. While bond lengths and
angles mostly tend to be fixed, torsional angles can vary widely, giving each protein its
structure. For a peptide with M residues, there are 3(M-1) torsional angles, with three
torsional angles (φ, ψ, and ψ) for every pair of residues. While φ and ψ torsional angles
have been extensively studied, relatively less is known about the ω torsional angle, the
one governing the geometry of the peptide bond. When making structural models, the
peptide bond is assumed to be planar, specifically, in a trans-like conformation, unless
known to be otherwise. With no general experimental method to distinguish cis- and
trans-peptide bonds, this is a serious shortcoming of protein structure determination.
This study is aimed at describing a solid-state NMR spectroscopic experiment
which is sensitive to the conformation about the peptide bond. Section 1.2.8 describes the
fundamental principle behind such a study, which involves correlating two anisotropic
interactions under magic-angle spinning. Such correlations are known to be sensitive to
the relative orientation of the two interactions. Chapter 3 describes the specific
experiments chosen for this study, with simulation results showing that it would be
possible for such experiments to differentiate between model compounds with trans and
80
cis peptide bonds. Chapter 2 has described the preparation of the model compounds with
high purity. Trans-peptide bond, realized with linear diglycine, was synthesized by solidphase peptide synthesis using standard protocols. Cis-peptide bond, realized with 2,5diketopiperazine, was synthesized using non-conventional microwave-assisted synthesis.
Both of these compounds yielded spectra with reliable signal-to-noise, and produced
REDOR decay patterns that agree well with simulation, as shown in Chapter 4.
`
Preliminary results from the simulations show that the primary objective, to
distinguish cis-peptide bonds from their trans counterparts, should be possible with the
experiment described here. However, more work is necessary to achieve the farther goal,
that of measuring deviation of a peptide bond from planarity. Simulations show this
experiment to be insensitive to such small changes in the ω-torsional angle.
Modifications are required to make the experiment suitable for this purpose.
The immediate next step would be to extend the applicability of this experiment
to a full protein. Potential systems to be studied by this method could be Barstar or
Acquaporin, each of which have a native cis-peptide bond, and are known to give good
quality solid-state NMR spectrum. In such a scenario, the experiment has to be modified
to include an additional chemical shift evolution dimension on 15N. This would result in a
2D
13
C-15N spectrum, where each cross-peak is modulated by information about the
geometry about the peptide bond. A series of such 2D spectra can be collected, and the
trajectory of each peak can be fit to simulations to extract the geometry information about
each peptide plane.
Additionally, in such a scenario, the experiment needs to be compensated for the
effects of side-chains, which were not present in the model compounds. Specifically, the
81
POST-C7 sequence is known to be sensitive to presence of
13
C in the side chains. A
potential solution could be introducing SEASHORE49 delays in between POST-C7
blocks.
Another potential area of improvement is to increase the experiment sensitivity,
so as to detect small changes in the ω-torsional angle. Primarily, sensitivity around ωangle of 180º is desired, as most peptide bonds exist in the trans-like conformation. One
option might be the use of carbonyl CSA instead of 13C-13C dipolar coupling. Preliminary
simulations and experiments (not shown) show that carbonyl CSA decays the
magnetization much faster (~ 0.2 ms) than the
13
C-15N dipolar coupling evolves (~ 1.5
ms). This problem may potentially be solved by using 15N-1H dipolar coupling instead of
13
C-15N dipolar coupling. 15N-1H dipolar coupling, reintroduced by T-MREV39, is known
to decay the magnetization in ~ 0.5 ms (Figure 3.3, taken from reference 39). Correlated
evolution of these two interactions, in a ratio to be determined by simulations, could be
expected to yield a more sensitive version of this experiment.
82
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