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Theses and Dissertations
2012
Applications of ion mobility mass spectrometry :
screening for SUMOylation and other posttranslational modifications
Quentin Dumont
The University of Toledo
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Dumont, Quentin, "Applications of ion mobility mass spectrometry : screening for SUMOylation and other post-translational
modifications" (2012). Theses and Dissertations. 306.
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A Thesis
entitled
Applications of Ion Mobility Mass Spectrometry - Screening for SUMOylation and
Other Post-Translational Modifications
by
Quentin Dumont
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Master of Science Degree in Chemistry
_________________________________________
Dr. Wendell P. Griffith, Committee Chair
_________________________________________
Dr. Timothy C. Mueser, Committee Member
_________________________________________
Dr. Max O. Funk, Committee Member
_________________________________________
Dr. Patricia R. Komuniecki,
Dean College of Graduate Studies
The University of Toledo
December 2012
Copyright 2012 Quentin Dumont ©
This document is copyrighted material. Under copyright law, no parts of this document
may be reproduced without the expressed permission of the author.
An Abstract of
Screening for SUMOylation and Other Post-Translational Modifications – Applications
of Ion Mobility Mass Spectrometry
by
Quentin Dumont
Submitted to the Graduate Faculty as partial fulfillment of the requirements for the
Master of Science Degree in Chemistry
The University of Toledo
December 2012
Post-translational modifications (PTMs) of proteins are fundamental processes
that trigger, regulate and terminate most of the cellular mechanisms by covalent
attachment of chemical moieties to substrate proteins. Conjugation to the Small
Ubiquitin-like MOdifier (SUMO) is a highly conserved and regulated posttranslational
modification that is often restricted to highly specific cellular events and a number of
obligatory nuclear regulatory processes. Although much is known about the processes of
SUMOylation and deSUMOylation, exactly how it mediates its various functions and 4
different human isoforms remains unclear. This is due in part to the extremely low
abundance of modified proteins (about 2% of a given substrate will become modified),
and the large expense of time and effort required to identify and characterize these
SUMOylated proteins by current methods.
This thesis presents the development and application of an ion mobility mass
spectrometry (IMMS)-based method of screening for isopeptides from SUMOylated
substrates. The model conjugates poly-SUMO2 and poly-SUMO3 were digested by
trypsin/chymotrypsin to produce small linear peptides and isopeptides with a QQQTGG
iii
tag on the substrate lysine. Using solution conditions to promote higher charge states and
IMMS, mass spectra for the larger isopeptides with +3 charge state were extracted from
the mass mobility plot. These isopeptides were confirmed using tandem mass
spectrometry. Interestingly, a neutral loss of 17 Da was observed for all isopeptides,
which resulted from the loss of ammonia due the acid-catalyzed rearrangement of the Nterminal glutamine residue of the isopeptide tag. The method was also applied to in vitro
SUMOylated RanGAP1 and Sp100 fragments, two known SUMO substrates.
IMMS was also applied to the identification of tyrosine-sulfation in peptides using
an approach that was developed for phosphorylated peptides. Analysis of a trypsin digest
of phosphorylated a-casein showed the expected decrease in drift time. A similar
reduction in drift time was observed for tyrosine sulfated synthetic peptide
DY*MGWMDF-NH2 (Spp) relative to its unmodified analog (DSpp). This decrease in
drift time was a result of the modified peptides becoming more compact due to
electrostatic interaction between the negatively charged modification and the positively
charged N-terminus. The tyrosine-sulfated peptide and its unmodified analog DSpp were
spiked in equimolar amount into a digest of bovine serum albumin. The band
corresponding to the modified peptide was clearly identified in the much more complex
mass mobility plot with a significant shift in the drift time for the tyrosine-sulfated
peptide.
IMMS is a highly promising method for analysis of PTMs due to its sensitivity
and selectivity. In this work we demonstrated its utility in screening protein digests for
isopeptides from SUMO-conjugated substrates and tyrosine-sulfated peptides using
model analytes. Future work will involve testing the limits of these methods including
peptide sequence, peptide size and detection limits in more complex cellular matrices.
iv
Acknowledgements
I would like to express my deepest appreciation to my advisor Dr. Wendell P.
Griffith for his constant help, his guidance and limitless patience. Thank you for teaching
me so much about mass spectrometry, for the excellent balance between your
encouragements when the morale was low and your criticism when I was losing focus. I
could not have wished for a better advisor! I would also like to thank my committee
members, Dr. Funk and Dr. Mueser for their helpful suggestions, and staying onboard
with me despite my complete change of project. I am grateful to the Chemistry
Department staff, for making my time here so smooth; and Leif Hansen who always
helped me with the MALDI when I needed it. I want to acknowledge Dr. Anderson, Dr.
Bryant-Friedrich, Dr. Lind, Dr. Funk, and Dr. Ronning for sharing their knowledge
during their amazing classes. I have learned so much! I also want to give a special thanks
to Edith Kippenhan, who is probably the best TA advisor on Earth!
I would like to thank my past and present labmates: Jingshu, David, Steven,
Camille, Anthony, Mallory and the grad students from other labs for the wonderful time
spent here; it was fantastic to meet all of these amazing people. A special thanks goes to
Lucile, who was always there for me, and with whom I have been on the craziest trips of
my life! I am thankful to my family and friends in France, for their unconditional love
and support. Thank you all for this unforgettable experience!
v
Table of Contents
Abstract ………………………………………………………………………………… iii
Acknowledgements ............................................................................................................ v
Table of Contents ..............................................................................................................vi
List of Figures .................................................................................................................. xii
List of Abbreviations ......................................................................................................xvi
Introduction ........................................................................................................................ 1
1.1. From Genes to Proteins: a Multi-Step Process With Increased Complexity ............ 1
1.1.1. Protein Synthesis: General Process ................................................................................ 1
1.1.2. The Diversification of the Proteome .............................................................................. 2
1.2. Post-Translational Modifications: Diversity and Functions ...................................... 4
1.2.1. The Different Types of PTMs ........................................................................................ 4
1.2.2. The Effect of PTMs on the Protein Substrates ............................................................... 5
1.3. Project Goal ............................................................................................................... 7
1.4. Organization of the Thesis ........................................................................................ 9
Mass Spectrometry and its Role in Proteomics ............................................................. 11
2.1. Principles of Mass Spectrometry............................................................................. 11
2.1.1. Fundamentals ............................................................................................................... 11
2.1.2. The Development of MS for Proteomics ..................................................................... 12
vi
2.2. Instrument Parts....................................................................................................... 13
2.2.1. Ionization Sources ........................................................................................................ 14
2.2.2. Mass Analyzers ............................................................................................................ 18
2.3. Synapt HDMS Instrument: Components and Features ........................................... 22
2.3.1. Intrument Parts ............................................................................................................. 22
2.3.2. Experimental Approaches ............................................................................................ 24
2.4. Applications of MS to Proteomics .......................................................................... 28
2.4.1. Definition of Proteomics .............................................................................................. 28
2.4.2. Protein Identity, Quantity and Structural Features ....................................................... 30
2.4.3. PTMs and Proteomics .................................................................................................. 32
Characterization of SUMOylated proteins by MS ....................................................... 34
3.1. Ubiquitin and SUMO .............................................................................................. 34
3.1.1. Structure and Functions of Ub ..................................................................................... 34
3.1.2. An Ubiquitin-Like Modifier: SUMO ........................................................................... 36
3.2. Existing Methods for the Detection of Ubls by MS ................................................ 40
3.2.1. Methods for Ubiquitin .................................................................................................. 40
3.2.2. Yeast Analog ................................................................................................................ 43
3.2.3. Mutagenesis ................................................................................................................. 44
3.2.4. Use of High-end MS Instrumentation .......................................................................... 45
3.2.5. Software Tools ............................................................................................................. 46
3.2.6. Griffith/Cotter: Tailored-Proteolysis Approach ........................................................... 48
3.3. Requirements for a New Method ............................................................................ 51
vii
Material and methods ...................................................................................................... 53
4.1. Materials .................................................................................................................. 53
4.2. In vitro SUMOylation of Proteins ........................................................................... 54
4.2. SDS-PAGE Gels ..................................................................................................... 54
4.3. Enzymatic Digestion ............................................................................................... 55
4.4. Zip Tip Procedures .................................................................................................. 56
4.4.1. Manufacturer Protocol ................................................................................................. 56
4.4.2. Preparation of Oligonucleotides Procedure ................................................................. 57
4.5. Mass Spectrometry .................................................................................................. 57
4.5.1. Nano-ESI-TOF/IM MS ................................................................................................ 57
4.5.3. MALDI MS .................................................................................................................. 58
4.6. In silico Digests ....................................................................................................... 59
4.7. Spiking of BSA Digests .......................................................................................... 59
Pre-Screening Method for SUMOylated Proteins ........................................................ 60
5.1. Proof-of-Concept for the Screening Method ........................................................... 60
5.1.1. Overview ...................................................................................................................... 60
5.1.2. ESI-MS Analysis of the Poly-SUMO-2 Digest ........................................................... 61
5.1.3. Ion Mobility Analysis of the Poly-SUMO-2 Digest .................................................... 63
5.1.4. Other SUMO Isoforms ................................................................................................. 69
5.2. Applications of the Method ..................................................................................... 70
5.2.1. In vitro SUMOylation: Sp100 and RanGAP1 .............................................................. 70
Analysis of Tyrosine O-Sulfation by IMMS .................................................................. 74
viii
6.1. Physiological Functions of Small PTMs ................................................................. 74
6.1.1. Sulfation ....................................................................................................................... 74
6.1.2. Phosphorylation ........................................................................................................... 76
6.1.3. Sulfation, Phosphorylation: Similarities and Differences ............................................ 77
6.2. Project Goal ............................................................................................................. 79
6.3. Optimization of the IMMS Experiments: Phosphorylation .................................... 80
6.3.1. Model System .............................................................................................................. 80
6.3.2. Results .......................................................................................................................... 81
6.4. Screening for Sulfated Peptides .............................................................................. 85
6.4.1. Model System .............................................................................................................. 85
6.4.2. Analysis of Spp and DSpp ........................................................................................... 85
6.4.3. Spiking of a Bovine Serum Albumin Digest with Spp ................................................ 88
6.4.4. The effect of positive versus negative ionization mode ....................................... 90
6.5. State of the Project and Conclusion ........................................................................ 94
Conclusions and Future Directions ................................................................................ 96
7.1. Conclusions ............................................................................................................. 96
7.2. Future Directions ..................................................................................................... 98
References ....................................................................................................................... 100
Appendix A: Tandem MS of the linear SUMO-2 peptides ........................................ 111
Appendix B: Tandem MS simulated distribution of SUMO-2 isopeptides .............. 116
Appendix C: ESI-MS and MALDI-MS data for Sp100 and RanGAP1 fragments . 119
Appendix D: -casein peptides identified .................................................................... 123
ix
Appendix E: BSA peptides identified ........................................................................... 127
x
List of Tables
1.1
Known protein post-translational modifications of amino acids side chains.......... 5
1.2
Common PTMs: the change in mass on the substrate and their functions ............. 7
4.1
Protein and reagent amounts and final concentrations used for in vitro SUMO
conjugation reactions ............................................................................................ 54
4.2
Chemicals and amounts used for the preparation of 15% acrylamide SDS-PAGE
gel .......................................................................................................................... 55
5.1
List of the major linear peptides detected from the poly-SUMO-2 dual digest.... 63
5.2
List of the SUMO isopeptides detected using ion mobility mass spectrometry ... 68
6.1
Comparison of some biological and biochemical features of phosyphorylation and
sulfation as PTMs; together with their properties during MS analysis ................. 79
D.1
List of the major linear peptides detected matching the masses of an in silico list
of -casein digest peptides.................................................................................. 123
E.1
List of the major linear peptides detected matching the masses of an in silico list
ofBSA digest peptides ........................................................................................ 127
xi
List of Figures
2-1
Schematic of a typical mass spectrometer ............................................................ 14
2-2
Formation of ions in ESI ....................................................................................... 16
2-3
Schematics of linear and single-stage reflectron TOF analyzers .......................... 21
2-4
Schematic of the Synapt HDMS system ............................................................... 23
2-5
The peaks found in MS spectra provide both qualitative and quantitative
information about the species being analyzed ...................................................... 25
2-6
Schematic of two data representations used for IMMS: ion mobilogram and drift
ion chromatogram ................................................................................................. 26
3-1
Scheme of the isopeptide bond formation during Ub or SUMO conjugation ...... 35
3-2
Crystal structures of human SUMO-1 and Ub ...................................................... 37
3-3
Sequence alignment of the C-terminal region of Ub and several Ubls ................. 42
3-4
Sequence alignment of the human SUMO isoforms C-terminal regions ............. 49
3-5
Schematic of an isopeptide carrying the 6 amino acid residues SUMO tag obtained
by dual digestion with trypsin and chymotrypsin ................................................. 50
3-6
Scheme of the N-terminal glutamine rearrangement into pyroglutamate ............. 51
5-1
Sequence of human SUMO-2 ............................................................................... 61
5-2
ESI-TOF MS mass spectrum of the poly-SUMO-2 dual digest ........................... 62
5-3
Mass-moblity plot of the trypsin/chymotrypsin digest of poly-SUMO-2 ............ 64
xii
5-4
Drift time total ion chromatogram of peptides detected from the
trypsin/chymotrypsin proteolysis of poly-SUMO2 .............................................. 65
5-5
Reconstructed mass spectrum of the z ≥ +3 peptides from the poly-SUMO-2 dual
digest ..................................................................................................................... 67
5-6
Comparison of the experimental and simulated isotopic distributions for peptide 2
of Table 5.2 ........................................................................................................... 67
5-7
MS/MS spectrum of the isopeptide 731.74 m/z (+3) (3 in Table 5.2) .................. 69
5-8
Reconstructed mass spectrum of the z ≥ +3 peptides from the poly-SUMO-2 dual
digest ..................................................................................................................... 70
5-9
Sequences of the commercial RanGAP1 and Sp100 fragments ........................... 71
5-10
SDS-PAGE of in vitro SUMOylation Sp100 and RanGAP1 fragments in presence
or absence of SUMO-1 and ATP .......................................................................... 72
6-1
ESI-TOF MS spectrum of an -casein trypsin digest ........................................... 82
6-2
Tandem mass spectrum of the precursor ion at m/z 1660.70 corresponding to the
singly phosphorylated peptide VPQLEIVPNpSAEER ........................................ 83
6-3
Mass-mobility plot of the -casein trypsin digest ................................................ 84
6-4
IMMS analysis of a solution of Spp in denaturing solution ................................. 87
6-5
MS/MS spectrum of the 1063.41 m/z (+1) peptide ............................................... 87
6-6
Positive ESI-TOF MS spectrum of a 5 M BSA trypsin digest spiked with an
equimolar ratio of Spp .......................................................................................... 89
6-7
Positive IMMS of a 5 M BSA digest spiked with a 1:1 molar ratio of Spp ....... 90
6-8
Analysis in positive and negative ionization mode of Spp solution ..................... 91
xiii
6-9
Negative ESI-TOF MS spectrum of a 5 M BSA trypsin digest spiked with an
equimolar ratio of Spp .......................................................................................... 92
6-10
MS/MS spectrum of the 1140.81 m/z (-1) peptide ................................................ 93
6-11
Negative IMMS of a 5 mM BSA digest spiked with a 1:1 molar ratio of Spp ..... 94
A-1
MS/MS spectrum of the linear SUMO-2 peptide 535.26 (+2) ........................... 111
A-2
MS/MS spectrum of the linear SUMO-2 peptide 599.35 (+2) ........................... 112
A-3
MS/MS spectrum of the linear SUMO-2 peptide 617.84 (+2) ........................... 112
A-4
MS/MS spectrum of the linear SUMO-2 peptide 749.87 (+2) ........................... 113
A-5
MS/MS spectrum of the linear SUMO-2 peptide 823.41 (+2) ........................... 113
A-6
MS/MS spectrum of the linear SUMO-2 peptide 901.46 (+2) ........................... 114
A-7
MS/MS spectrum of the linear SUMO-2 peptide 1106.58 (+1) ......................... 114
A-8
MS/MS spectrum of the linear SUMO-2 peptide 1342.57 (+1) ......................... 115
B-1
MS/MS spectrum of isopeptide 2 in Table 5.2 ................................................... 116
B-2
Comparison of the experimental and simulated isotopic distributions for peptide 1
of Table 5.2 ..................................................................................................................... 117
B-3
Comparison of the experimental and simulated isotopic distributions for peptide 3
of Table 5.2 ..................................................................................................................... 117
B-4
Comparison of the experimental and simulated isotopic distributions for peptide 4
of Table 5.2 ..................................................................................................................... 118
C-1
ESI-TOF MS spectrum of the in vitro SUMOylation of Sp100 fragment by
SUMO-1 .......................................................................................................................... 119
C-2
MALDI-TOF MS of the trypsin digested Sp100 fragment................................. 120
xiv
C-3
ESI-TOF MS spectrum of the in vitro SUMOylation of RanGAP1 fragment by
SUMO-1 .......................................................................................................................... 121
C-4
MALDI-TOF MS of the trypsin digested RanGAP1 fragment .......................... 122
D-1
MS/MS spectrum of the -casein peptide 615.32 (+1) ...................................... 123
D-2
MS/MS spectrum of the -casein peptide 748.37 (+1) ..................................... 123
D-3
MS/MS spectrum of the -casein peptide 1267.69 (+1) .................................... 124
D-4
MS/MS spectrum of the -casein peptide 1384.73 (+1) .................................... 124
D-5
MS/MS spectrum of the -casein peptide 1759.90 (+1) .................................... 125
E-1
MS/MS spectrum of the BSA peptide 517.34 (+1)............................................. 126
E-2
MS/MS spectrum of the BSA peptide 545.38 (+1)............................................. 128
E-3
MS/MS spectrum of the BSA peptide 649.38 (+1)............................................. 129
E-4
MS/MS spectrum of the BSA peptide 660.40 (+1)............................................. 120
E-5
MS/MS spectrum of the BSA peptide 689.43 (+1)............................................. 130
E-6
MS/MS spectrum of the BSA peptide 712.42 (+1)............................................. 130
E-7
MS/MS spectrum of the BSA peptide 789.53 (+1)............................................. 131
E-8
MS/MS spectrum of the BSA peptide 922.56 (+1)............................................. 131
E-9
MS/MS spectrum of the BSA peptide 927.57 (+1)............................................. 132
E-10
MS/MS spectrum of the BSA peptide 974.53 (+1)............................................. 132
E-11
MS/MS spectrum of the BSA peptide 1014.71 (+1)........................................... 133
E-12
MS/MS spectrum of the BSA peptide 1163.73 (+1)........................................... 133
E-13
MS/MS spectrum of the BSA peptide 1305.82 (+1)........................................... 134
E-14
MS/MS spectrum of the BSA peptide 1479.93 (+1)........................................... 134
E-15
MS/MS spectrum of the BSA peptide 1567.89 (+1)........................................... 135
xv
List of Abbreviations
ACN .......................... Acetonitrile
AmAc ........................ Ammonium Acetate
AmBic ....................... Ammonium Bicarbonate
AMP .......................... Adenosine Monophosphate
AP ............................. Atmospheric Pressure
ATG8 ........................ Autophagy-Related Protein 8
ATG12 ...................... Autophagy-Related Protein 12
ATP ........................... Adenosine Triphosphate
BSA ........................... Bovine Serum Albumin
CE ............................. Collision Energy
CENP-C .................... Human Centromer Protein C
CHCA ....................... -Cyano-4-Hydroxycinnamic Acid
CI............................... Chemical Ionization
CID ............................ Collision Induced Dissociation
DNA .......................... Deoxyribonucleic Acid
DSpp ......................... Desulfated Peptide
DTT ........................... Dithiothreitol
E. coli ........................ Escherichia coli
E1 .............................. Ubiquitin-Activating Enzyme
E2 .............................. Ubiquitin-Conjugating Enzyme
E3 .............................. Ubiquitin Ligase
ECD........................... Electron Capture Dissociation
ESI............................. Electrospray Ionization
ETD ........................... Electron Transfer Dissociation
FAB ........................... Fast Atom Bombardment
FT-ICR ...................... Fourier Transform Ion Cyclotron Resonance
FUBI ......................... Fau and its Ubiquitin-Like Domain
GTP ........................... Guanosine Triophosphate
HDMS ....................... High-Definition Mass Spectrometry
HEPES ...................... 4-(2-hydroxyethyl)-1-Piperazineethanesulfonic
xvi
HGP........................... Human Genome Project
HIV ........................... Human Immunodeficiency Virus
HPLC ........................ High Performance Liquid Chromatography
HUB1 ........................ Histone Mono-Ubiquitination Protein 1
IKB ......................... Kappa Light Polypeptide Gene Enhancer in B-Cell Inhibitor, Alpha
IMMS ........................ Ion Mobility Mass Spectrometry
ISG15 ........................ Interferon-Stimulated Gene 15
IT ............................... Ion Trap
LC ............................. Liquid Chromatography
LDI ............................ Laser Desorption/Ionization
MALDI ..................... Maxtrix-Assisted Laser Desorption/Ionization
mRNA ....................... Messenger Ribonucleic Acid
MS ............................. Mass Spectrometry
MS/MS ...................... Tandem Mass Spectrometry
NEDD8 ..................... Neural Precursor Cell Expressed Protein, Developmentally DownRegulated 8
PDB ........................... Protein Database
PTM .......................... Post-translational Modification
Q ................................ Quadrupole
RanGAP1 .................. Ran GTP-ase Activating Protein 1
RF .............................. Radiofrequency
RNA .......................... Ribonucleic Acid
RP .............................. Reverse Phase
S. cerevisiae .............. Saccharomyces cerevisiae
SDS-PAGE ............... Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis
SILAC ....................... Stable Isotope Labeling by Amino Acids in Cell Culture
SILIS ......................... Stable Isotope-Labeled Internal Standard
SPITC ........................ Sulfophenyl Isothiocyanate
Spp ............................ Sulfated Peptide
SRM .......................... Single Reaction Monitoring
SUMO ....................... Small Ubiquitin-Like Modifier
TEAA ........................ Triethylammonium Acetate
TFA ........................... Trifluoro Acetic Acid
TOF ........................... Time-of-Flight
TPST ......................... Tyrosylprotein Sulfotransferase
T-wave ...................... Traveling Wave
TWIMS ..................... Traveling Wave Ion Mobility Spectrometry
xvii
Ub .............................. Ubiquitin
Ubl............................. Ubiquitin-Like Modifier
UCRP ........................ Ubiquitin Cross-Reactive Protein
UFM1 ........................ Ubiquitin-Fold Modifier 1
URM1 ....................... Ubiquitin-Related Modifier 1
UV ............................. Ultraviolet
xviii
List of Symbols
cm .............................. Centimeter
m ............................. Micrometer
Da .............................. Dalton
eV .............................. Electron Volt
mM ............................ Millimolar
M ............................. Micromolar
m/z ............................. Mass-to-Charge Ratio
min ............................ Minute
Th .............................. Thomson
xix
Chapter 1: Introduction
1.1. From Genes to Proteins: a Multi-Step Process With Increased
Complexity
1.1.1. Protein Synthesis: General Process
All living organisms are maintained and regulated by mechanisms involving
proteins, the building blocks of any living species. Each protein carries out a specific
function, and is encoded by genes present in deoxyribonucleic acid (DNA). As the
complexity of an organism increases, the number of proteins required also increases.
Because the synthesis of so many proteins would demand a lot of energy, living
organisms have developed mechanisms to change the function of proteins after synthesis:
post-translational modifications (PTMs). Post-translational modification is the covalent
modification of a protein by a chemical moiety in order to change its function, thus a
single protein can be involved in different mechanisms depending on its modification
state. Most PTMs are reversible, which allows for proteome diversity. The understanding
of all biochemical mechanisms and pathways in the body cannot be achieved without the
study of PTMs. As the current methods available for the analysis of PTMs are not always
optimal, the work presented in this thesis is aimed at the development of new mass
spectrometry-based methods for the screening of select PTMs.
1
Proteins are linear polymers of amino acids, chemical subunits linked to each
other by peptide bonds. The range of weights, structures and functions for proteins is
extremely wide, and together with polysaccharides and nucleic acids, two other types of
biological macromolecules, proteins are involved in every single process occurring in the
cells. The wide variety of protein functions ranges from catalyzing biological reactions1
to helping fold other proteins.2, 3 Due to their omnipresence in any organism, their proper
biosynthesis is inherent to all life. Each protein has a specific amino acid sequence, which
is defined by the nucleotide sequence of the gene encoding the protein.4
1.1.2. The Diversification of the Proteome
One gene with a unique sequence has the capacity to encode only one protein of a
given amino acid sequence. The number of genes in an organism would define the
number of proteins and consequently, the complexity of the proteome (set of proteins
expressed by an organism). Following the revolution of the Human Genome Project
(HGP),5 the genome of 46 organisms has been sequenced. Interestingly, D. melanogaster,
a species of drosophila, was found to have about 26,000 genes, while humans possess a
genome of approximately 30,000 genes. Moreover, it was discovered that the genes in
humans are able to produce more than 1,000,000 different molecular protein species,
representing a complexity about two orders of magnitude higher than if one gene encoded
only one protein.6 The comparison of the complexity of both organisms was the first clue
that led to the conclusion that proteome diversity is not directly related to the number of
genes.
2
Indeed, it was later discovered that there are several mechanisms to generate high
diversity in the proteome. Several processes occur before or during protein synthesis.
They include mRNA splicing7 and editing,8 and alternative promoter.9 The last pathway,
which will be the focus of this thesis, that explains the much increased complexity of the
proteome compared to the genome is post-translational modification. PTM is the
chemical modification of a protein that occurs shortly after biosynthesis (once the RNA
has been translated) or at any point in its lifespan. PTMs are not directly encoded by
genes but can modify the function, location and turnover of a single protein. The
attachment of various biochemical functional groups (such as acetyl, lipid, carbohydrate,
and sulfate) extends the range of functions of the protein by changing the chemical nature
of the modified amino acid, or inducing structural change (for instance, formation of
disulfide bridges). The wide variety of PTM reactions are catalyzed by an equally
important number of enzymes, and can occur at one or several places either on the side
chain of amino acids constituting the primary sequence or on the backbone of the protein
itself. Although PTMs have been identified in prokaryotes, the extent of modifications is
much higher in eukaryotes both in terms of diversity and occurrence.6 It has to be noted
here that the modification of a protein is typically not homogenous: because of alternative
splicing and a combination of various modifications described above, a single gene can
lead to multiple gene products. Consequently, the amount of protein in a given
modification state is minute compared to the total amount of gene products.
3
1.2. Post-Translational Modifications: Diversity and Functions
1.2.1. The Different Types of PTMs
PTMs can be divided into two main groups: the ones that covalently modify the
substrate protein by addition of a chemical moiety or polypeptide chain, and others that
cleave the backbone at a specific peptide bond determined by the primary sequence. For
the purpose of this dissertation, most of the discussion is focused on the first kind of
PTM, leading to covalent modification of the protein.
Of the 20 common amino acids produced by human cells, only 15 of them can
undergo post-translational modification of their side chain. These amino acids and some
of their known modifications are listed in Table 1.1. The combination of the 15
modifiable amino acids and the variety of modifications possible for each residue leads to
an extremely broad range of possible PTMs possible for a single protein. Each covalent
modification has a specific effect on the modified protein, whether it affects the formal
charge or the overall protein structure. The extent of change can vary greatly, for example
N-methylation adds 14 Daltons (Da) to the protein mass, while addition of ubiquitin
increases the total mass by more than 8,000 Da.
4
Table 1.1: Known protein modifications of amino acids side chains6
Residue
Mechanisms
Arg
N-methylation, N-ADP-ribosylation
Asn
N-methylation, N-ADP-ribosylation
Asp
Phosphorylation
Cys
S-hydroxylation, disulfide bond formation, phosphorylation, S-acetylation
Gln
Transglutamination
Glu
Methylation, carboxylation, polyglycination, polyglutamination
Gly
C-hydroxylation
His
Phosphorylation
Lys
N-methylation, N-acetylation, Ubiquitination, SUMOylation
Met
Oxidation to sulfoxide
Pro
C-hydroxylation
Ser
Phosphorylation
Thr
Phosphorylation, O-glycosylation
Trp
C-mannosylation
Tyr
Phosphorylation, sulfation, ortho-nitration
1.2.2. The Effect of PTMs on the Protein Substrates
PTMs are of great importance in most cellular pathways and processes. When a
protein gets modified, one or several of its features is changed. As an example,
acetylation of a positively charged  amine group on a lysine residue side chain will
neutralize its positive charge. Phosphorylation of a serine residue increases negative
charge and hydrophilicity, thereby altering the protein’s properties and in some cases
structure may be altered. Intra-molecular interactions can be disrupted or modified. Those
modifications ultimately have an effect on the function of the protein. Enzymes have very
5
specific active sites due to the interactions created between the binding pocket and the
substrate. If an amino acid gets modified and triggers a change in the structure of the
active site, even a small one, the enzyme can undergo a complete loss of its catalytic
activity, and the pathway in which it is involved is then down-regulated. A list of some
common and important PTMs, mass changes and their functions is provided in Table 1.2.
Due to the variety of the possible modifications and the broad range of possible chemical
and physical changes to a protein, PTMs are involved in the regulation of most cellular
events, and play an important role in all steps of the proteins life cycle. For example,
poly-ubiquitination of short-lived proteins results in their degradation by the 26S
proteasome.10 Comprehensive knowledge of PTMs is a requirement for complete
understanding of the mechanisms taking place in the cell. Some modifications are
currently very-well characterized (for example phosphorylation) and their analysis is
carried out by routine experiments. However, understanding of mechanisms involving
larger PTMs (SUMO, in particular) has been hindered by the lack of analytical methods
providing fast and reliable results.
6
Table 1.2: Common PTMs: the change in mass on the substrate and their functions11
PTM type
Phosphorylation
pTyr
pSer, pThr
Mass
change
(Da)
+ 42
Methylation
Acetylation, fatty acid
modification
Farnesyl
Myristoyl
Palmitoyl
Glycosylation
N-linked
O-linked
+ 14
Cellular localization and signals targeting,
membrane tethering, mediator of proteinprotein interactions
+ 204
+ 210
+ 238
Excreted proteins, cell-cell
recognition/signaling, reversible, regulatory
functions
Glycosylphosphatidylinositol (GPI) anchor.
Membrane tethering of enzymes and
receptors, mainly to outer leaflet of plasma
membrane
Protein stability and protein-ligand
interactions
Modulator of protein-protein and receptorligand interactions
Intra- and intermolecular crosslink, protein
stability
Possible regulator of protein-ligand and
protein-protein interactions, also a common
chemical artifact
Protein stability, blocked N-terminus
Destruction signal
Oxidative damage during inflammation
> 800
203, > 800
> 1,000
Hydroxyproline
+ 16
Sulfation (sTyr)
+ 80
Disulfide bond formation
-2
Deamidation
+1
Pyroglutamic acid
Ubiquitination
Nitration of Tyr
Reversible, activation/inactivation of
enzymes activity, modulation of molecular
interactions, signaling
Protein stability, protection of N-terminus.
Regulation of protein-DNA interactions
Regulation of gene expression
+ 80
+ 80
Acetylation
GPI anchor
Function and notes
- 17
> 1,000
+ 45
1.3. Project Goal
Post-translational modifications are a crucial aspect to the understanding of cells
as they trigger, regulate and terminate most of the mechanisms that take place. Studying
the proteome without taking PTMs into account would give an incomplete, if not
7
incorrect, picture of the function of each protein. There is consequently a real need for the
development of fast and reliable methods for the detection and characterization of PTMs.
This goal has been hindered in the past by the fact that the occurrence of PTMs is usually
small (for instance, SUMOylation happens for only 2% of substrate proteins)12 and a
number of PTMs are labile. Both of these issues make their analysis more challenging
and requiring instrumentation and methods with very low detection limit, sensitivity, and
capability for conserving labile modifications.
SUMO is an ubiquitin-like modifier that has gained importance in the scientific
community over the past few years. While ubiquitin function is quite well known,
especially its role in protein degradation by the proteasome,13 SUMO has only recently
become a subject of interest. The implication of SUMOylation in many nuclear pathways,
together with finding that the deregulation of SUMOylation leads to various diseases,
have increased current interest in this modification. To gain a better understanding,
effective techniques must be available for the analysis of SUMOylation. Although some
research groups have developed reliable MS methods for the analysis of PTMs, and more
specifically SUMOylation (see Chapter 3 for more details), most of them lack
physiological relevance and/or are highly time-consuming.
One of the goals of the research carried out in the Griffith laboratory is to develop
new mass spectrometry-based methods for the characterization of analytes that have
proven too challenging for other currently available analytical methods. In that
perspective, the goal of the project presented in this thesis is to develop and apply
techniques using the existing MS instrumentation for the reliable, efficient, and accurate
screening of PTMs, in particular SUMO. The ion mobility mass spectrometry utility of
8
the Waters Synapt High Definition Mass Spectrometry (HDMS) instrument was used for
all of the work presented here. Due to its ability to separate ions, based on molecular
shape, size and net charge in addition to m/z, IMMS is a convenient technique for the
analysis of PTMs. Applications of IMMS to the analysis of SUMOylation and tyrosine Osulfation are presented in this thesis.
1.4. Organization of the Thesis
The experiments carried out for the completion of this research involved used of
techniques including, but not limited to, in-solution enzymatic digests, in vitro
modification of proteins, and desalting using ZipTips. The mass spectrometers that were
used included the Bruker Esquire-LC (ESI-QIT MS), Bruker Daltonics UltrafleXtreme
(MALDI-TOF/TOF MS), and the Waters Synapt HDMS (ESI Q-TOF MS) equipped with
nano-ESI source and traveling wave ion mobility MS.
This thesis has been divided into seven chapters, each addressing a specific aspect
of the project. Firstly, an overview of the mechanisms that increase proteome complexity
is provided, with focus on the diversity and the functions of post-translational
modifications.
In this research, PTMs were analyzed using mass spectrometry, a
technique that has become the method of choice for the analysis of biological samples.
Consequently, the second chapter provides insight into the field of mass spectrometry
with its developments, the features of each part of a mass spectrometer and the intricate
relation between MS and proteomics. Chapter 3 provides an overview of the
SUMOylation reaction, together with the existing MS based methods that are used to
detect and identify SUMOylated proteins. The goal of this thesis research was to develop
9
a new method that can rapidly analyze post-translational modified proteins with fewer
purification steps. The material, methods and instrumentation used for these experiments
are presented in Chapter 4. The first project was the development of a technique using ion
mobility mass spectrometry as a tool to allow fast and simple screening for SUMOylated
isopeptides. The proof-of-concept details and applications of this method are highlighted
in Chapter 5. Chapter 6 focuses on similar IMMS approaches for screening and
identification of tyrosine O-sulfation sites in proteins. The thesis concludes with an
overview of the results and presentation of some possible future directions.
10
Chapter 2: Mass Spectrometry and its Role in
Proteomics
2.1. Principles of Mass Spectrometry
Of all the currently used analytical techniques, very few are as versatile as mass
spectrometry (MS). Owing to the many developments in MS instrumentation over the last
few decades, MS provides unequaled performance in terms of sensitivity, detection limits,
ease of use and speed of analysis. The applications are very diverse and range from
nanoparticle analysis to the study of intact ribosomes. The development of electrospray
ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI) makes MS
suitable for analysis of biological samples and currently MS experiments are widely used
for protein and biomolecules analysis.
2.1.1. Fundamentals
During a mass spectrometry experiment, ions are separated in the instrument
based on their mass-to-charge ratios and detected in proportion to their abundance.14 The
collected data are plotted in a mass spectrum, where the abscissa is a scale of mass-tocharge ratio (m/z, or Thomson, Th)15 and the ordinate usually relative intensity. The yaxis may also display ion count in some data representations. In order to be detected, the
11
analytes have to be gas phase ions. In ESI-MS analysis, typically the solution containing
the analyte is introduced into the instrument via an ionization source. As the name
suggests, this part of the instrument is used to vaporize and ionize the analytes. After
desolvation, ions are focused and sorted in the mass analyzer, which discriminates the
ions depending on their m/z. They finally hit the detector which counts the number of ions
for each m/z value and sends an electrical signal interpreted by the computer.16
Because the ions formed are short-lived and unstable, the instrument is under high
vacuum (usually from 10-3 to 10-6 torr pressure) to prevent degradation of the ions. The
source is sometimes at atmospheric pressure, and a gradient of pressures (called
differential pumping) makes the transition between atmospheric pressure (AP) and high
vacuum. Low pressure ensures a longer mean free path, which is the distance an ion can
travel without colliding into another particle. It is necessary for the instrument to have a
large mean free path so that the ions can reach the detector without undergoing collisions,
which can alter the path of the ion.
2.1.2. The Development of MS for Proteomics
The first mass spectrometer, a “parabola spectrograph” was built in 1913 by
Thomson.17 However, it took about a century of developments and breakthroughs in the
field of MS for mass spectrometers to be suitable to the analysis of large biological
samples. Although a wide variety of ionization techniques were developed quite early
(electron impact, chemical ionization, etc.), none of them could keep the analyte intact,
i.e. without fragmentation. In the mid-1980s, a true need for analysis of full-length
proteins with high molecular masses emerged, and the techniques available at that time
12
did not meet the criteria for the desired analysis.18 Although ionization by Fast-Atom
Bombardment (FAB)19 could prevent most of the fragmentation of proteins, this
technique was limited by the fact that it produced mainly singly-charged ions. Proteins
with a molecular weight above 1,000 Da could not be measured by the mass analyzers
available.
The invention of ESI, the first “soft” ionization method, by John Fenn in 198920
overcame all of these problems. Briefly, ESI creates multiply charged species, keeping
the analyte without any fragmentation, which enables analysis of large proteins. Indeed,
because of the multiple charging of analytes, the m/z values are in a range that can be
measured by common mass analyzers. The importance of this discovery that changed the
field of MS was rewarded by a portion of the Nobel Prize in Chemistry in 2002. More
details about ESI are provided in the Section 2.2.1. Some other soft ionization techniques,
notably MALDI, developed by Tanaka in 1988,21 have been used for the analysis of
biological samples, but none can compete with the ease of use and efficiency of the ESI.
Owing to the advances made in the past, MS has grown increasingly important in the field
of proteomics and is currently one of the most versatile and efficient method for gaining
knowledge about proteins.
2.2. Instrument Parts
All MS instruments have the same basic “building blocks”: an ionization source, one or
several mass analyzers, and a detector, (Fig. 2-1). Although for most instruments the
ionization source is found under vacuum, atmospheric pressure sources have also been
developed; they include electrospray ionization, atmospheric pressure matrix-assisted
13
laser desorption/ionization and atmospheric pressure chemical ionization (AP-CI). The
choice of each part is extremely important, as their properties can be very different; thus
the choice of each component of the instrument is application-driven.
Figure 2-1: Schematic of a typical Mass Spectrometer.
2.2.1. Ionization Sources
A large number of ionization sources are currently available, each one suitable for
a range of applications. They can be classified into two groups: the organic ionization
sources used to ionize organic and biological compounds; and the inorganic sources that
are more suitable for studying metals, oxides, etc. For the purpose of this thesis, the focus
will be on two organic ionization sources, widely applied to the analysis of biological
samples: ESI and MALDI.
14
2.2.1.1. Electrospray Ionization
Electrospray ionization is one of the most widely used techniques in the field of
proteomics due to its unique properties. ESI produces highly-charged analytes while
preventing them from fragmentation as they are being ionized. Although it is currently
used for many different families of compounds, such as small organic molecules, etc.,
ESI was developed for proteins and it remains the best ionization source for the analysis
of biological samples.
Typically, a solution of the analyte (millimolar, mM concentration or lower) in a
polar volatile solvent is injected via a syringe into a metal capillary at a low flow rate (120 L/min). A high voltage, 2-6 kV, is applied to the tip of the capillary, creating an
electric field between the capillary and the counter electrode located at the entrance of the
instrument, 1 or 2 centimeters (cm) away from the tip. As the solution exits the capillary,
the strong field induces a charge accumulation at the liquid surface at the end of the tip,
resulting in the formation of a Taylor cone, followed by dispersion into an aerosol
containing highly charged droplets. As the solvent is evaporated, a point is reached when
the surface tension (which holds the droplet together) is equal to the columbic repulsion
known as the Rayleigh limit. As the solvent continues to evaporate, the charges become
too close to one another, and the droplet “explodes” into smaller ones. This process is
repeated until a totally desolvated gas-phase ion is obtained. A schematic of the formation
of charged ions in ESI is presented in Fig. 2-2. It is interesting to note that despite the fact
that ESI is so widely used, the exact mechanism is currently not fully understood.
Nevertheless, some studies are leading the way to the better understanding of the physical
and chemical processes involved in ESI.22, 23
15
Figure 2-2: Formation of ions in ESI. The blue dots represent the analyte molecules. a:
solvent evaporation. b: “Coulombic explosion”. Figure adapted from24
The Synapt HDMS that was used in this work is equipped with a nanoESI source,
with a lower flow rate of about 0.5 to 1 L/min. The spray needle is replaced by a
borosilicate glass capillary of a few L volume with a diameter at the tip of 1-4 m. The
initial droplets produced by the nano-ESI are much smaller compared to conventional ESI
(about 200 nm diameter vs. 20-200 m). The lower flow rate affords one greatly reduced
sample consumption, facilitates improved desolvation and consequently sensitivity.
As ESI produces analytes that are multiply charged species, the charge state
distributions in mass spectra provide much information about the studied compounds. For
instance, it has been shown that proteins that are denatured give mass spectra displaying
peaks at higher charges, with higher intensities and with a wider charge state distribution
than when they are analyzed in their native, more folded conformation.25, 26 A denatured
protein has more solvent-exposed surface area and is statistically more likely to
16
accommodate a larger number of protons than more folded conformers thereby. Due to all
of the advantageous features mentioned above, the range of applications for the ESI is
very broad. ESI-MS-based experiments using ESI are further described in Section 2.4.
2.2.1.2. MALDI
Matrix-Assisted Laser Desorption/Ionization was developed by Karas and
Hillenkamp in 198727 and shortly thereafter was applied to the analysis of proteins with a
molecular weight over 100 kDa by Tanaka.21 MALDI was developed as an improvement
to the already existing Laser Desorption/Ionization (LDI) technique, where the analyte is
fixed on a metal plate and irradiated with an ultraviolet (UV) laser. However this
ionization source had several limitations: extensive fragmentation due to the large amount
of energy absorbed by the sample, low sensitivity, poor reproducibility and very high
dependence of the signal on the capacity of the analyte to absorb UV light. MALDI is
based on the same principle, except that the analyte is co-crystallized with an excess of
matrix, an organic molecule chosen to absorb light and enhance ionization of the sample.
The matrix molecules are typically aromatic to absorb UV light, acidic to donate protons
to the analyte, and easy to co-crystallize with the analyte of interest.28, 29 The addition of
the matrix makes the ionization of the analyte indirect. Thus, for each laser pulse the
sample is not fragmented as efficient desorption/ionization occurs.30 In addition, using a
matrix enables the spot where the laser hits to be refreshed, greatly increasing the shot-toshot reproducibility of the analysis.
MALDI has many interesting features that make it suitable for the analysis of
large biopolymers and to the field of proteomics. The main one is that unlike ESI,
MALDI mainly produces singly charged ions. It becomes convenient for analysis of
17
complex peptide mixtures, as one peptide produces only one peak in the resultant mass
spectrum, while for ESI each peptide produces multiple charged species, greatly
increasing the number of peaks and the complexity of data analysis. For larger analytes
like proteins, the +1 charge state peak is oftentimes the most intense, but it not
uncommon for the higher charge states to be detected as well. Another advantage of using
MALDI is its relative tolerance for common ionization suppressors in protein and peptide
samples such as salt, glycerol and urea. Such compounds can be present at a low
concentrations in biological samples and not greatly interfere with the analysis.31 This
enables faster determination of the analytes as a perfect purification is not required.
MALDI MS is a very sensitive technique, requiring only nano- to fentomoles of sample.
It has to be pointed out that MALDI MS is not suitable for compounds with a low
molecular mass. Matrix and fragment ions produce high-intensity peaks in the range m/z
≤ 500. Small analytes, which give peaks in the same area of the mass spectrum, are likely
to not be detected or overlap with the matrix peaks. In a typical MALDI analysis, the
minimum m/z is often set at 400 in order to exclude most of the matrix peaks. Pulsed
sources like MALDI work really well with time-of-flight mass analyzers.
2.2.2. Mass Analyzers
The improvements in the field of MS are not only due to the development of
ionization sources, mass analyzers are just as important to get a reliable analysis with
quality data. Mass analyzers are the part of MS instruments that separates ions based on
their m/z. There are several types of mass analyzers, each one possessing a specific m/z
range, resolving power and mechanism of ion transmission. Two main classes can be
distinguished: the scanning mass analyzers which allow only ions with a specific m/z to
18
go through (e.g. quadrupole), and the simultaneous transmission mass analyzers, for
which all of the ions pass through at the same time (e.g. time-of-flight).
2.2.2.1. Quadrupole
The single quadrupole (Q) analyzer is a device composed of four metallic rods
perfectly parallel to each other. Two are connected by a constant voltage, and the other
two rods by a radiofrequency voltage. The polarity of each rod switches from positive to
negative in a cyclic fashion. Each pair of rods is 180° out of phase with the other. The
principle of the quadrupole is that the rods create an oscillating electric field within which
only ions with a given m/z are stable.32 When a positive ion enters the quadrupole, it is
attracted by the negative rod due to electrostatic interactions. If the rod switches to
positive before the ions reaches it, the latter is repulsed from the rod and keeps moving
towards the exit end of the mass analyzer. Otherwise, it crashes into the rod, becomes
neutral and is not detected. The modulation of the currents (intensity, frequency of
oscillation, etc.) enables scanning for a large range of m/z values. The advantage of using
scanning analyzers, such as a quadrupole, is that as only ions with very close m/z are
transmitted at the same time and reach the detector, the sensitivity is greatly improved.
The detector has to scan only a small range of values, making the detection more
efficient.
Quadrupoles are inexpensive, long-lived and commonly used. However, they are
limited by an upper m/z of 4,000; they are not suitable for analysis of large compounds
(especially proteins), unless the ionization source produces multiply charged ions. Thus,
ESI-Q is a widely used combination for the analysis of proteins, while MALDI-Q is
seldom employed. Quadrupoles, when not used as mass analyzers, can also serve as ion
19
guide at the entrance of the mass spectrometers, in order to focus the ions and ensure a
stable trajectory. The first commercial quadrupole mass analyzer was developed by
Finnigan in 1994.33
2.2.2.2. Time-of-flight
Although the principles of time-of-flight (TOF) mass analyzers were described in
1946,34 the first design for a linear TOF analyzer was only published a decade later in
1955 by Wiley and McLaren,35 later becoming the first commercial TOF instrument.
The separation of ions in a TOF analyzer is based on the fact that ions with
different m/z have different velocities, so they take different amounts of time to travel a
defined given distance: ions are dispersed in time. By measuring the time needed for each
ion to reach the detector, one can determine its m/z value and obtain a mass spectrum for
the sample. Practically, the ions produced as packets, are initially accelerated by an
electric field. They then enter the flight tube (a field-free region) where they are separated
based on their velocities before reaching a detector situated at the extremity of the tube
(in linear mode). It is important to note that all the ions entering the drift tube have the
same kinetic energy. There are two types of TOF mass analyzers: linear and reflectron.36
The design differences are shown in Fig. 2-3.
20
Figure 2-3: Schematics of linear (top) and single-stage reflectron (bottom) TOF
analyzers. Adapted from Cotter et al. Anal. Chem. 199937
In reflectron mode, an electrostatic reflector creates a retarding field that deflects
the incoming ions back into the drift tube. The reflectron is situated opposite the ion
source (about the same place as the detector in linear mode) while the orthogonal detector
is off-axis with respect to the initial ion beam so that it can be placed co-axially to the ion
source. Ions with higher velocity go deeper into the reflectron field, while slower ions do
not spend as much time in it. Hence, reflectron mode enables a larger flight distance,
significantly improving the resolution. However, the main advantage of the reflectron is
to correct the dispersion in the initial kinetic energies for ions with the same m/z leaving
the source, so they all end up reaching the detector at the same time. This is illustrated on
Fig. 2-3 (bottom), where three ions of same m/z, regardless of whether or not their kinetic
21
energy is lower (eV – U0) or higher (eV + U0), have their trajectories corrected by the
reflectron and are detected simultaneously. This prevents peak broadening and improves
the resolution of the data obtained. Despite these advantages and the dramatic
improvement in resolving power, there are some drawbacks to using reflectron mode. The
main drawback is the limitation in the m/z range that can be analyzed. While in linear
mode, there is virtually no limitation for the molecular weight of the analyte (masses
above 300 kDa have been detected with MALDI-TOF MS,38, 39 the reflectron is limited to
≤ 10,000 m/z. TOF analyzers have a high transmission efficiency (i.e. the number of ions
exiting the mass analyzer is almost the same as the number of ions entering it) leading to
higher sensitivity. Due to the longer distance the ions have to travel, the sensitivity in
reflectron mode is usually lower than in linear mode. TOF mass analyzers provide an
excellent synergy with pulsed ionization techniques that produce packets of ions, such as
MALDI.
2.3. Synapt HDMS Instrument: Components and Features
2.3.1. Intrument Parts
The instrument that was used for the majority of the work presented in this thesis
is the Synapt HDMS system (Waters Corp.), a hybrid quadrupole ion mobility separation
orthogonal acceleration time-of-flight mass spectrometer equipped with a nanospray
source (Fig. 2-4).
22
(e)
(a)
(b)
(c)
(d)
Figure 2-4: Schematic of the Synapt HDMS (Waters Corp.)40
The nanospray ionization source (a), so-called “Z-shaped” source is engineered to
effectively get rid of solvent molecules and neutral species before they can enter into the
instrument. Pushers repel the charged ions to the entrance while the unaffected neutral
molecules are pumped away by the vacuum system. Once inside the instrument, the ion
beam is guided by a traveling-wave (T-wave) ion guide (b), whose role is to transfer ions
from the pressurized ion source region to high-vacuum. Next, the quadrupole (c) acts as a
mass filter, which stabilizes the trajectory of the ions. A focusing lens focuses the ions
before they enter the tri-wave region (d) of the instrument. The tri-wave region is
comprised of three T-waves connected in tandem. The T-wave cell consists of a stackedring radio frequency (RF) ion guide which incorporates a repeating sequence of transient
voltages applied to the ring electrodes. These voltage pulses create a traveling electric
field that propels ions through the background gas present in the mobility cell. For a given
23
charge state, ion-neutral collision frequency increases with extended ion conformation
resulting in higher propensity to roll back over the waves. The time it takes for an ion to
drift through the cell depends on its mobility; the wave period and height; as well as the
gas pressure. Ions with high mobility (compact shape) are better able to keep up with
traveling waves and are pushed more quickly through the cell. Ions with low mobility
(extended structures) crest over the waves more often and have to wait for subsequent
waves to push them forward, resulting in longer drift times.41
In the Synapt HDMS, each cell of the tri-wave region has a specific function. The
trap T-wave ensures high-efficiency by trapping the ions and increasing their number. It
can also be used as a collision cell to fragment ions of interest. The IMS T-wave cell is
where the ion mobility separation occurs: ions are discriminated based on their charge,
size and shape. Ion mobility MS is described more thoroughly in thesis Section 2.3.2.3.
Finally, the packets of ions exiting the transfer T-wave are repelled into the TOF mass
analyzer (e). The TOF can be operated in either of two modes: “V-mode”, for which the
ions are reflected twice before reaching the detector; or “W-mode” where the resolution is
increased dramatically as the ions travel almost twice the distance, but this improvement
is at the cost of sensitivity.
2.3.2. Experimental Approaches
2.3.2.1. MS Analysis
The most basic MS experiment that can be carried out is a survey scan. Basically
the sample is introduced into the mass spectrometer and ionized; the ions are separated by
the mass analyzer and cataloged according to their m/z; and finally their intensities
24
measured and plotted on a mass spectrum. Although it appears to be a simple
experimental scheme, a great deal of information can be extracted from a typical mass
spectrum. The peaks displayed on mass spectra provide both qualitative (position on the
m/z scale) and quantitative (peak height or area) information about the analyte, as seen in
Fig. 2-5. However, it has to be pointed out that the peak height is not enough to achieve
quantitation without proper standards as different analytes can have different ionization
efficiencies leading to varying intensities, even if they were introduced in equimolar
amounts. Because any slight change in the analyte conformation, mass or environment
triggers a change in the detected peaks, even a simple MS scan under suitable
experimental conditions is powerful enough to analyze non-covalent complexes, for
instance. Usually, MS scans are the first step of any MS-based experimental workflow as
they provide an overview of the sample’s contents and complexity.
Figure 2-5: The peaks found in MS spectra provide both qualitative and quantitative
information about the species being analyzed.
25
2.3.2.2. MS/MS Analysis
Tandem mass spectrometry (MS/MS) experiments involve at least two stages of
mass analysis, in conjunction with a dissociation process. Commonly, two mass analyzers
are required for this method: the first one selects an ion of given m/z (chosen by the user),
which then undergoes spontaneous or activated fragmentation, leading to product ions
and neutral fragments. Activation can be achieved in multiple ways, commonly using
collisions with neutral gas molecules42 or a solid material,43 electrons44,
45
or photons.46
The daughter ions are separated by a second mass analyzer, detected, and the mass
spectrum for the fragments is obtained. In the case of the Synapt HDMS system, the
selection of the parent ion m/z is achieved by the quadrupole. Selected ions are then
fragmented by violent collision with inert gas atoms (Argon) in the trap T-wave cell; the
fragments are then mass analyzed by the TOF analyzer and detected.
The interest of MS/MS lies in its ability to break down a complex molecule to its
basic structural features. Some practical applications of MS/MS are structure elucidation
(small organic molecules, differentiation of isomers and stereoisomers), selective
detection of target compound class, ion-molecule reaction studies, and protein
sequencing.47
2.3.2.3. IMMS
Ion mobility spectrometry is a way of separating ions based on their interactions
with an inert buffer gas (usually He or N2)48 as they fly through a drift tube. When
coupled with mass spectrometry, this technique is referred to as ion mobility mass
26
spectrometry49 and becomes a powerful analytical tool capable of separating isomers,
isobars and conformers; reducing chemical noise and measuring ion sizes (cross section).
The strength of this approach relies on the ability of IMMS to differentiate between
isomeric and isobaric analytes that have the same molecular mass and nominal mass,
respectively. IMS can be carried out in four different ways depending on the type of
instrument: drift-time IMS, aspiration IMS, field-asymmetric waveform IMS; and
traveling wave IMS (TWIMS, which is used in the Synapt HDMS). The mode of
operation of TWIMS was described in Section 2.3.1. of this thesis. It is also operated at
reduced pressure, which is a unique characteristic compared to the other IM
spectrometers. TWIMS exhibits high transmission efficiency and separative power
although the resolution is not as good as in conventional drift time methods.50
IM is extremely efficient when coupled with TOF mass analyzers as this
technology provides an analyzer which is capable of providing full mass spectra on
timescales short enough to enable profiling of millisecond wide ion mobility peaks.51 The
data obtained when carrying out an IMMS experiment are presented as a mass-mobility
plot (ion mobilogram), representing the mass-to-charge ratio against the drift time (or
vice versa). A unique property of IMMS spectra is that in ion mobilograms, regardless of
how they are plotted, mass-mobility correlations (commonly called “trend-lines”) are
observed for classes of ions. Any statistically relevant deviation from these trend lines by
an ion gives information regarding its structural compactness. Another way of presenting
the data is to use a drift time ion chromatogram, where the intensity (or ion count) is
plotted against the drift time; this representation is used to compare the drift times of all
the ions detected. A schematic of the data obtained in IMMS is presented in Fig. 2-6.
27
A
B
Figure 2-6: Two common data representations used for IMMS: ion mobilogram or massmobility plot (A) and drift time total ion chromatogram (B). The dashed lines represent
the mass mobility correlations and go through all the ions with a same charge state.
There is an almost infinite number of applications for IMMS, some of them
include inorganic chemistry (metabolic profiling of inorganic ions),52 gas-phase ion
structure studies, isomer separation in complex mixtures, analysis of saccharides,
peptides, proteins, nucleic acids, drugs and metabolites. The importance of IMMS has
grown increasingly in the field of proteomics due to the rapid and high-resolution
separation it provides.53
2.4. Applications of MS to Proteomics
2.4.1. Definition of Proteomics
Characterization of proteins present in a biological system (proteome) provides
insight into the function and complexity of that system. The proteome is not only
28
complex, but also spatially, temporally and chemically dynamic. Proteomics is the
systematic study of all of the proteins expressed by a genome, cell, tissue or organism at a
given time point under defined conditions. The term appeared in print for the first time in
1995.54 Proteomics aims to answer questions such as: what proteins are expressed in a
cell, and in which amounts; how these expression levels relate to function? How do
proteins interact with each other? How do PTMs help to regulate the function of proteins?
Two approaches have been developed: global proteomics, which experiments are
designed to characterize as many proteins expressed by a genome as possible; and
targeted proteomics for which the number of proteins analyzed is much lower (e.g. only
proteins that are phosphorylated). Historically, proteomic experiments were mainly
conducted using 2D gel electrophoresis for protein separation but the progress made in
analytical instrumentation extended the range of techniques used; for example cell
imaging, array and genetic readout experiments. However, MS has grown increasingly
important in proteomics as the development of the instrumentation enabled the analysis of
complex protein samples,55 and currently most of the proteomics efforts are MS-based.
As previously explained in Chapter 1, the proteome is much more complex than
the genome, which makes proteomics a more challenging field compared to genomics.
Indeed, different cells express different proteins, or a same protein can be expressed by
two different cells but not at the same level. Localization of the protein, interactions with
other biomolecules, PTMs and conformational changes in protein structures are some of
the obstacles that highlight the difficulty of proteomics. Practically, in order to overcome
these challenges, proteomics experiments involve a step to reduce the complexity of the
29
representative proteome sample, analysis by MS and bioinformatics tools for fast data
processing.
2.4.2. Protein Identity, Quantity and Structural Features
One aspect of proteomics, usually the first carried out on an unknown system, is
protein characterization. Identifying the proteins comprising the proteome of a given cell
or organism is a necessary step before conducting more advanced experiments. Two main
experimental methods have been developed: the “bottom-up” and “top-down”
approaches.
2.4.2.1. Bottom-up Approach
The most common MS-based proteomics technique to study proteins is the
bottom-up approach. The workflow is as follows: a sample containing several proteins is
isolated and enzymatically digested into peptides; trypsin is commonly used as the
positive ions created by cleavage C-terminally to lysine and arginine residues enhance
ionization efficiency and detection. The resultant peptide mixture is fractionated using
liquid chromatography (LC) or other methods (affinity purification among them). This
step allows for enrichment of the sample and reduces the number of peptides entering the
mass spectrometer at any given time, thereby increasing sensitivity. As the peptides elute
from the chromatography column (typically reverse phase), they are infused directly into
the MS and ionized by the ESI source. At that point, a complete scan is taken and the
peptide masses are compared against in silico digested protein sequences from databases
of fully-sequenced proteins (Mascot, Sequest, etc.).56 If more information is needed for a
30
given peptide, tandem mass spectrometry can be carried out, and the MS/MS data leads to
its amino acid sequence.
The bottom-up approach is very popular because of its many advantages: this
method is sensitive; samples can be analyzed with a high-throughput as some steps can be
automated, and software for data interpretation is available. This approach is fast and
reliable and has been applied to many systems: the shotgun method allowed for
identification of 42 subunits of the mitochondrial complex I, which represents 95% of the
complex.57 It was also used to analyze MCF-7 cell lines expressing the zinc-finger or the
proline-rich domain of retinoblastoma-interacting-zinc-finger protein,58 and identification
of a single mutation in Escherichia coli (E. coli) RNA polymerase was achieved
following this workflow.59 In the case of PTMs, the bottom-up approach is interesting as
it produces peptides that carry the modification and can be distinguished by mass from
the unmodified peptides (see Thesis Section 2.4.3.). Despite these many advantages, the
efficiency of this technique decreases dramatically as the complexity of the sample
increases.
2.4.2.2. The Top-Down Approach
The top-down proteomics approach refers to the analysis of intact proteins, which
are not enzymatically digested prior to the MS experiment. Bottom-up and top-down
approaches each have specific advantages and drawbacks that have been thoroughly
discussed in the past.60, 61 The top-down approach has the advantage of being less timeconsuming and to provide the mass of the intact protein and can consequently highlight
modifications, cleavages or isoforms of a given protein. However, analyzing intact
proteins is more challenging as the ionization is limited and the detection of high
31
molecular weight compounds is reduced. Oftentimes, the full-length protein is fragmented
and the masses of the obtained fragments can be compared against database values.
The top-down approach is very efficient for large proteins or proteins expressed at
high levels. It has been applied to the analysis of intact membrane proteins.62 Soft
ionization MS preserved the covalent structure of the proteins that were then fragmented;
the sequence of fragments define the original native covalent state of the protein. Proteins
with masses greater than 200 kDa were successfully identified by this technique using
electrospray additives, heated vaporization, and separate non-covalent and covalent bond
dissociation.63 For the characterization of PTMs, top-down is a more suitable approach as
the fragment ion mass data are much more specific than the masses of the peptides from
protein’s digest. This has been demonstrated by Sze and co-workers in 2002 as they
reported the characterization of PTMs within one residue for a 29 kDa protein.64
2.4.3. PTMs and Proteomics
Qualitative MS-based proteomics analyses of PTMs on purified proteins are
usually achieved by peptide mapping. The protein is digested with one or more
proteolytic enzymes in order to cover as much of its sequence as possible. Protein
modifications are then identified by correlation of the measured masses and sequence
information derived from MS/MS data. Peptides carrying a modification display a shift in
mass corresponding to the weight of the attached moiety, and are usually easily
identifiable. This can be carried out manually or with the use of a number of available
software packages including Mascot, Sequest, etc.65 This untargeted approach is
commonly used because a single protein can be multiply modified, and it is generally
32
faster to screen for all PTMs instead of targeting them one by one.66 The detection of
modifications on histones, proteins that are heavily and variably modified, is an example
of successful methods for PTM analysis by proteomics.67
In addition to this general method, several techniques for targeted identification of
modifications have been developed.68 The experimental design for large-scale analysis of
PTMs always involves some sort of sample enrichment, usually tandem affinity
purification.69 For instance, the efficiency of the sample enrichment in phosphopeptides
using titanium oxide (TiO2) columns was beneficial to the analysis of phosphorylation in
proteins. Phosphorylation is one of the modifications associated with biological
regulation as it is involved in many mechanisms.70, 71, 72
Small stable modifications such as methylation and acetylation are relatively easy
to detect and characterize by MS, given that they only change the molecular weight of the
protein by a few Daltons. However, some PTMs add much larger moieties to the
substrate. Ubiquitin (Ub) is a protein that links to the  amino group of a lysine residue
and adds 8.5 kDa to the protein. The Small Ubiquitin-Like Modifier (SUMO) is an 11
kDa protein that modifies its substrate the same way as ubiquitin. The great diversity of
these PTMs requires equally as diverse analytical methods for their analysis.
33
Chapter 3: Characterization of SUMOylated proteins
by MS
3.1. Ubiquitin and SUMO
3.1.1. Structure and Functions of Ub
Ubiquitin is regulatory protein found in almost all tissues of eukaryotes.
Interestingly, ubiquitin is one of the most highly conserved proteins among eukaryotic
organisms, but totally absent from the superkingdoms Eubacteria and Archaea.13
Ubiquitin is a 76 amino acid protein with a molecular weight of 8564.84 Da. The human
Ub was isolated and sequenced by Edman degradation for the first time in 1975.73 The
protein is now very well-known and fully characterized; it is also one of the first PTMs to
be identified.74
Substrate proteins are covalently modified by Ub through an enzyme cascade
requiring the concerted action of three enzymes referred as E1, E2 and E3.75 Firstly, E1
(ubiquitin-activating enzyme) activates Ub through an ATP-dependent process by
producing an Ub-adenylate intermediate. Ub is then transferred to the E1 active site
cysteine residue with loss of adenosine monophosphate (AMP) and formation of a
thioester bond between the Ub C-terminus and the E1 cysteine sulfhydryl group. Through
34
a trans(thio)esterification reaction, Ub is transferred to E2 (ubiquitin-conjugating
enzyme). Finally, with the assistance of one of hundreds of E3 enzymes (ubiquitin ligase),
the last step of the process results in the formation of an isopeptide bond between the Cterminal glycine of Ub and the -amino group of the target lysine (Fig. 3-1).76
R
1
NH
O
H
C
CH
CH
O
C
NH
NH2
H
OH
CH2
H2C
C-terminal di-glycine
H2C
R
2
CH
NH
Target lysine side chain
CH2
O
C
R
3
- H2 O
O
R
1
NH
H
C
CH
CH
O
NH
C
H
isopeptide bond
NH
CH2
H2C
CH2
H2C
R
2
NH
CH
O
C
R
3
Figure 3-1: Scheme of the isopeptide bond formation during Ub or SUMO conjugation to
a target lysine.
35
One specific feature of this modification is that Ub can modify itself and form
chain or branched structures with various lengths and linkages, providing versatile means
of cellular regulation. Interestingly, the various Ub branched and chain structures have
different effects on the target protein, thus this single modification displays various
functions. It has been shown that Ub is involved in the mechanisms regulating cell
development, growth, and apoptosis and signal transduction processes;77 however it is
mainly known for triggering degradation of proteins by the 26S proteasomes through
ATP-dependent attachment of a specific polymeric chain of Ub.78, 79
3.1.2. An Ubiquitin-Like Modifier: SUMO
While the Ub sequence is highly conserved phylogenetically, it has been found
that by contrast, a number of proteins sharing a similar fold with Ub (but very little
sequence homology) are highly different from one organism to another. Despite low
identity to Ub, along with similar fold these ubiquitin-like modifiers (Ubls) share some
general features, such as the chemistry of modification and the enzyme cascade leading to
reversible and covalent binding. The currently known Ubls include: interferon-stimulated
gene 15 (ISG15); neural precursor cell expressed protein, developmentally downregulated 8 (NEDD8); autophagy-related protein 8 (ATG8); autophagy-related protein 12
(ATG12); fau and its ubiquitin-like domain (FUBI); ubiquitin-related modifier 1
(URM1); ubiquitin-fold modifier 1 (UFM1); histone mono-ubiquitination protein 1
(HUB1); ubiquitin cross-reactive protein (UCRP); and small ubiquitin-like modifier
(SUMO).80 These proteins all display the diglycine motif at their C-terminal (in mature
form), a necessary feature for covalent binding to the substrate proteins.
36
One Ubl in particular, the small ubiquitin-like modifier, an approximately 11 kDa
protein, has been shown to covalently modify a large number of proteins to regulate many
cellular processes including gene expression, chromatin structure, signal transduction and
maintenance of the genome.81,
82, 83
Contrary to what the name suggests, SUMO only
shares about 18% sequence homology with Ub but the both proteins share a -GRASP
fold, as illustrated in Fig. 3-2. (adapted from literature):84 the same secondary motifs are
found in both structures, although SUMO possesses an N-terminal extension absent from
the Ub protein. The core of both proteins is formed by four -sheets around an -helix. A
small helical structure is also present at the periphery of the proteins, with a high solvent
exposure.
Figure 3-2: Human SUMO-1 (left, PDB: 1AR5) and Ub (right, PDB: 1UBQ) share a
similar fold: they both display the same secondary motifs. SUMO-1 has a floppy Nterminal extension absent in the Ub structure.
37
The E1 and E2 enzymes involved in SUMOylation are closely related to those
involved in ubiquitination. However, some differences have been identified between the
two mechanisms: for example while many E2 enzymes have been identified for the Ub
process, only one conjugating enzyme, UbcH9, is currently known for SUMO.
Moreover, E3 is crucial for the specificity of Ub attachment, but does not appear to be
even necessary for SUMOylation to properly occur. The chemistry of modification and
isopeptide bond formation is the same as well. Because the mechanisms of SUMOylation
and ubiquitination are so similar and has been shown to target the same lysine residues in
certain proteins, it has been hypothesized that SUMO and Ub act as antagonists.84 For
instance, SUMOylation stabilizes the nuclear factor of kappa light polypeptide gene
enhancer in B-cells inhibitor, alpha (IkB) while ubiquitination leads to its degradation.
SUMOylation occurs on a consensus acceptor site consisting of the sequence KxE/D,
where  is a large hydrophobic amino acid, x can be any amino acid residue, and K is the
modified lysine.85 It has to be noted that this sequence is not an absolute rule, as it has
been reported that about 40% of proteins can be SUMOylated outside of this consensus
sequence, demonstrating differences in substrate specificity.12 Other studies have
identified two alternative consensus sequences for SUMOylation: one includes a
phosphorylation site (KxExxpSP where pS is a phosphoserine)86 and the other is a
negatively charged amino acid-dependent extended motif that enhances SUMOylation.87
Four SUMO isoforms have been identified in humans: SUMO-1, -2, -3 and -4.
Yeast cells express a simple SUMO analog, Smt3p, which has been widely used in the
early stages of SUMO research due to its easy expression in large amounts. Moreover, the
primary sequence of Smt3p allows for straightforward determination of modified peptides
38
by MS (see Thesis section 3.2.2. for more details). SUMO-2 and SUMO-3 are closely
related sharing 95% sequence homology, but only 50% sequence identity with SUMO1.88 SUMO-4 has a restricted expression pattern, with the highest levels found in the
kidneys.89 Some substrates can be modified by both SUMO-1 and -2/3, while Ran GTPase activating protein 1 (RanGAP1) for instance, the first substrate ever found for
SUMOylation,90 is almost exclusively modified by SUMO-1. Topoisomerase II is
predominantly a substrate for SUMO-2/3. It has been mentioned previously that the
function of Ub is dependent on the chain or branched structures formed on the substrate
protein. This feature is found to a lesser extent with SUMO: SUMO-2 and -3 can
covalently bind to themselves via the lysine residue located in a consensus sequence near
the N-terminus and form polymeric chains (not branched). SUMO-1 does not have a
consensus acceptor site, though it has been reported to form polymers in vitro. No proof
of SUMO-1 polymerization has yet been demonstrated in vivo, leading to the idea that
SUMO-1 would act as a chain terminator.12 The significance of SUMO polymer chains
has to be investigated for further understanding of the physiological functions of
SUMOylation.
SUMO has grown as a protein of interest, due to its implication in many cellular
(especially nuclear) mechanisms. Data obtained after deregulation of the SUMOylation
process
suggests
that
diseases
such
as
cancer,
pathogenic
infections
and
neurodegenerative disorders could be linked to malfunction of the SUMO machinery. It is
also important to understand SUMOylation in order to generate reliable cell models to
predict the toxicity of drugs in humans.91 Several approaches have been reported in the
past to characterize SUMO-modified proteins.
39
3.2. Existing Methods for the Detection of Ubls by MS
3.2.1. Methods for Ubiquitin
Historically, ubiquitination was the first PTM that was reported,74 and since then
its identification has become a routine procedure, especially with the development of
proteomics experiments involving MS. Because some MS-based methods to characterize
SUMOylated proteins are based on the principles used for Ub identification, an overview
of the technique is provided.
The early stages of identification and characterization of Ub substrates were
difficult due to low steady state levels and the presence of a varying number of Ub
molecules attached to different molecules of the same species. The first Ub identification
method was reported in 1985 by Haas and Bright;92 they conducted sodium dodecyl
sulfate polyacrylamide gel electrophoresis (SDS-PAGE) of whole cell extracts followed
by immunoblot analysis with anti-Ub antibody. They detected a complex profile of poorly
resolved high molecular mass proteins together with a few well revolved low molecular
mass proteins. This workflow was optimized and applied in 1992 by Beers and coworkers.93 Due to the inefficiency of the detection of Ub-modified proteins, which are
present in low amounts in the sample, methods involving specific affinity purification
were developed; pre-concentration of the sample in ubiquitinated proteins allowed for
detection and identification of multiple Ub substrates. Oftentimes, the studies of protein
ubiquitination involves three major steps: affinity purification, proteolytic digestion and
tandem MS. Proteins are expressed with an epitope tag genetically attached to the Nterminus of ubiquitin, which enables post-expression affinity purification.94 Peng et al.
reported a study in which Ub conjugates were identified by expression of His 6-tagged
40
proteins, Ni-affinity capture, trypsin digestion, chromatographic separation and MS/MS
analysis, leading to the discovery of 1075 Ub candidate substrates.95
Ubiquitination adds 8.5 kDa to the mass a protein, making its direct analysis by
MS challenging: the large size of the modification makes ionization of the substrate less
efficient; increases the complexity of the MS/MS spectra; and prevents the use of
bioinformatics tools and databases to process the data obtained. Thus, the use of
enzymatic digestion arose, making the MS analysis more straightforward. Most
frequently, trypsin is used: it cleaves the C-terminal peptide bond of lysine and arginine
residues, unless followed by a proline. The advantage of using this protease is that the
mature ubiquitin sequence terminates with an arginine residue followed by two glycines.
Thus, after trypsin digestion, the modified lysine residue contains a diglycine tag (+ 114
Da). While the unmodified peptides display a linear structure, the Ub-modified ones carry
a tag, making them branched. Because a peptide bond is formed between the substrate
lysine and the Ub glycine, they are referred to as isopeptides. The mass shift caused by
the – GG tag is easily identifiable by MS or MS/MS and can be used as a diagnostic tool
for the identification of modified peptides. Because of its small molecular weight, the
strength of this technique is the simplicity of the analysis once the diglycine tag is
obtained by digestion.
Due to the success of this method, the idea of applying it to the Ubls has arisen.
However, the primary sequence of many Ubls is such that the proteolytic cleavage site is
located several residues from the C-terminus, making the tag relatively long. Figure 3-3
presents the sequence alignment of the C-terminus of the four SUMO isoforms; human
Ub; the yeast analog of SUMO, Smt3p; and two other Ubls: UCRP and NEDD8. With the
41
exception of UCRP and NEDD8, the length of the tags resulting from trypsinolysis
(shown in bold font) for the SUMO proteins are greatly increased compared to Ub.
MS/MS data collected for isopeptides from the human SUMO isoforms would mostly
provide sequence information for the tag, which would not enable mapping of the
modified lysine or identification of the substrate protein. Due to this, a number of other
approaches have been developed.
SUMO-2_HUMAN
SUMO-3_HUMAN
SUMO-4_HUMAN
SUMO-1_HUMAN
SMT3_YEAST
UBIQ_HUMAN
UCRP_HUMAN
NEDD8_HUMAN
QIRFRFDGQP
QIRFRFDGQP
QIRFRFGGQP
SLRFLFEGQR
SLRFLYDGIR
QQRLIFAGKQ
LFWLTFEGKP
QQRLIYSGKQ
INETDTPAQL
INETDTPAQL
ISGTDKPAQL
IADNHTPKEL
IQADQTPEDL
LEDGRTLSDY
LEDQLPLGEY
MNDEKTAADY
EMEDEDTIDV
EMEDEDTIDV
EMEDEDTIDV
GMEEEDVIEV
DMEDNDIIEA
NIQKESTLHL
GLKPLSTVFM
KILGGSVLHL
FQQQTGG
FQQQTGG
FQQPTGG
YQEQTGG
HREQIGG
VLRLRGG
NLRLRGG
VLALRGG
Figure 3-3: Sequence alignment of the C-terminal region of Ub and several Ubls
showing the low conservation of primary sequence. Tags obtained on modified peptides
after trypsin digestion as shown in bold.
As mentioned previously, Ub is highly conserved across species, making the
method described above applicable for most organisms. However, Ubls have little
homology with Ub, translating into their C-terminal residues not necessarily giving a -GG
tag when cleaved with trypsin. It is true in particular for SUMO isoforms, which yield a
19 (SUMO-1) or 32 (all others) residue tag. The MS-based methods currently used for
analysis of Ubl-modified proteins include the study of the SUMO yeast analog; creation
of mutated SUMO proteins; high-end MS with FT-ICR instruments; the use of
specifically designed software; and a tailored-proteolysis approach.
42
3.2.2. Yeast Analog
The SUMO analog protein in Saccharomyces cerevisiae (S. cerevisiae) possesses
an arginine residue at the sixth position from the C-terminus. Proteolytic digestion with
trypsin consequently leads to a 5 residues tag (EQIGG), adding a supplementary 484.2 Da
to the mass of the substrate peptide.
The general procedure for analysis of SUMOylated substrates in S. cerevisiae
follows the same key steps as for Ub. The proteins are expressed with both His6 and
FLAG tags, and are purified by tandem affinity chromatography, with a nickel column
followed by a FLAG-affinity purification.96 The tandem purification has the advantage of
dramatically reducing the time spent sequencing non-SUMOylated proteins with MS/MS;
the time saved can be redirected to analyze the peptides of interest. Furthermore, using
two columns increases the confidence that the proteins isolated are actual targets of the
SUMO pathway. The purified and pre-concentrated proteins are then digested with
trypsin leading to two kinds of peptides: linear peptides (from SUMO and from target
protein), and isopeptides containing the modified lysine residue and SUMO tag. Modified
peptides feature a missed cleavage site at the modified lysine and an increased mass due
to the tag. Database searching finally enables for rapid interpretation of the tandem MS
data obtained.97 In some studies, mutagenesis was used to obtain a -GG tag and apply the
same method as for Ub.98
Yeast is an interesting model as it allows for straightforward genetic and
biochemical studies. Despite this important feature, the relevance of this method is
questionable regarding the goal of gaining a better understanding of SUMOylation to
43
endogenous human proteins. Characterizing proteins modified by Smt3 can point to some
interesting leads but is not sufficient by itself for the complete understanding of human
SUMO physiological targets and specific functions.
3.2.3. Mutagenesis
Site directed mutagenesis has been used to help reduce the size of the tag carried
by SUMO-modified peptides: by mutating a residue close to the C-terminus to an
arginine or a lysine, a trypsin cleavage site is created and the tag is reduced to only a few
amino acids. In 2005 Knuesel and co-workers reported a SUMO-1 mutant yielding a
diglycine tag after trypsinolysis.99 They observed that Ub and two other Ubls possess a
tryspin cleavage site (arginine) at the third residue away from the C-terminus; however,
SUMO-1, -2 and -3 have a threonine residue at the third position, preventing trypsin from
cleaving at this site. To mimic Ub, the mutation T95R was implemented. As Ub and
SUMO are so similar in their activity, it was assumed that the mutation would not induce
any changes in reactivity. This hypothesis was confirmed by their experiments. In 2010,
Blomster et al. reused the T95R SUMO-1 idea and added many other mutations (C52S,
H75K, V87K, V90C, and Q92C) on the C-terminus of SUMO-1 in order to purify the
protein based on cysteine-affinity columns.100 Using this short-tag mutant in combination
with MS/MS, they were able to identify the SUMOylation sites in twelve human proteins.
Although the T95R provides a trypsin cleavage site that allows for a short tag with trypsin
digestion, threonine and arginine do not share many similarities in terms of structure and
polarity. Using this observation, some groups have moved the mutation to an amino acid
of closer properties to arginine. The sixth residue away from the C-terminus of all human
SUMO isoforms is a glutamine. Mutating to an arginine residue induces little change in
44
the structure of the side chain, conserving the reactivity and functions of SUMO.101 The
five amino acid fragment obtained after trypsin digestion results in fewer fragment ions
from the isopeptide tags and facilitates data interpretation with common databases and
bioinformatics tools. Despite the advantages of using the mutagenic approach to SUMO
analysis, the main issue with such procedures is the amount of time and effort needed to
complete them.
3.2.4. Use of High-end MS Instrumentation
Digestion with trypsin produces extremely large isopeptides; the tag itself for
SUMO-2, -3 and -4 consists of up to 32 amino acid residues. Two main issues arise with
the MS analysis of such SUMOylated peptides. The first one is peaks are obtained from
both the long SUMO tag and the substrate peptides, significantly increasing the
complexity of the tandem mass spectra. Moreover, because the size of the tag is
oftentimes much larger than the substrate peptide, most of the fragments obtained
originate from the SUMO isopeptide tag and not the substrate peptide. This is a key issue
as the sequence information of the substrate peptide is crucial for mapping the modified
lysine and for identification of the modified protein. In order to solve these issues, highenergy fragmentation is required to increase the number of fragments and allow for
detection of substrate peaks. However, even common instruments such as Q-TOF mass
spectrometers, capable of high-energy fragmentation, do not produce enough fragments to
obtain the information needed.
A solution to this challenge is to use high-resolution and high-sensitivity MS
instruments such as the FT-ICR; despite the high cost, these mass spectrometers currently
45
have the best performance in terms of limit of detection, sensitivity and resolving power.
The use of non-ergodic (fast heating) fragmentation methods, especially electron capture
dissociation (ECD) and electron transfer dissociation (ETD), allows for direct bond
cleavage and dramatically increased fragmentation efficiency. Coupling FT-ICR
instruments with ETD/ECD fragmentation mechanisms can provide substrate peptide
information even when carrying a long SUMO tag. The use of FT-ICR MS for the
detection of ubiquitination sites was reported in 2004 by Cooper et al.102 They extended
this approach one year later to SUMOylation,103 achieving measurement of a SUMO1:RanGAP1418-587 intact conjugate with the mass accuracy of 2.7ppm. Moreover, the
digestion with trypsin of in vitro modified wild type proteins led to the mapping of the
substrate lysines. The efficiency of the FT-ICR also allowed for the characterization of
the number of SUMO chains when the protein was poly-SUMOylated, which is of
importance to understand how SUMOylation regulates the mechanisms in which
substrates are involved. This methodology was also successfully applied to the
identification of the modification sites of the human centromere protein (CENP-C).104
Despite the quality of the data that can be obtained by FT-ICR MS, the instrumentation
required is fairly expensive and not commonly found in many laboratories. Consequently,
this is not a good method for routine analysis with commonly available MS instruments.
3.2.5. Software Tools
For the analysis of small PTMs, such as tyrosine phosphorylation or lysine
methylation, the data collected with a LC/MS/MS method can be processed with the help
of a database. Standard database searching algorithms compare the experimental MS/MS
spectra to theoretical spectra generated in silico and identify any resulting mass matches.
46
Thus, based on the m/z and pattern of the obtained peaks, the primary sequence of the
peptide (modification included) can be identified. However, database searching is only
applicable to small modifications that do not fragment during analysis, and alter one of
the peptide fragment ions by a specific, indivisible mass. In the case of SUMOylation, the
fragmentation within the isopeptide tag produces peaks in the mass spectrum, which
obscure the pattern of the modified target peptide. Because several ion series are
overlapping on the same spectrum, database searching is inefficient for SUMOylation
data processing.
Some research groups have developed an automated approach for SUMO analysis
using software to process MS/MS datain. SUMmOn, developed by Pedrioli and coworkers, is an automated pattern recognition tool that detects diagnostic PTM ion
fragment series within complex collision-induced dissociation (CID) mass spectra to
identify modified peptides and modification sites within those peptides.105 SUMmOn is
based on an algorithm that extracts intensities for any user-defined b- and y-ion series
from every MS/MS scan in a specific analysis. Because SUMO is attached by its Cterminus, the y-ion series generated by the modification is dependent on the mass of the
substrate peptide. Consequently, the yn1+-ion series must be recalculated for each MS/MS
scan by subtracting the masses of the bn1+ (independent) fragments from the singly
charged precursor ion mass, and adding the mass of one hydrogen atom. At the end of the
procedure, SUMmOn calculates two scores: one for the modification and one for the
target peptide. The use of this software in combination with alternative proteases (LysC
among them) allows for maintaining of the identity of the original Ub and Ubl conjugate.
Indeed, if a target is modified by Ub and two other Ubls producing a -GG tag after
47
trypsinolysis, the identity of the modifier is lost after digestion. However, using this
approach in combination with LysC digestion and SUMmOn analysis allows for
comprehensive identification of Ub/Ubl-modified peptides. Furthermore, Jeram et al.
identified putative NEDDylation sites, and previously unpublished SUMO and NEDD8
chain topologies.106 A few years after the development of SUMmOn, “ChopNSpice”, a
simple and straightforward database tool was presented.107 The software relies on the idea
that MS/MS fragmentation of branched peptides is similar to the fragmentation of a linear
peptide with a miscleaved lysine residue and the SUMO peptide at its N-terminus.
However, databases do not include SUMO as a putative modification at lysine residues.
To address this problem, ChopNSpice automatically generates SUMO-modified FASTA
sequences of proteins in silico. These sequences are then implemented in a database
search (SEQUEST, MASCOT, etc.), to identify acceptor sites for SUMO conjugation.
This method is limited to fairly simple samples. Once the complexity of a sample
increases, the SUMmOn database increases as well, leading to a higher number of eligible
target peptides. To overcome this issue, the instrument must possess an excellent mass
resolution and high duty cycle in order to avoid peak crowding. The development of
software tools for the PTM identification is of great value for data interpretation but does
not bring anything new to the protein preparation or analysis by MS.
3.2.6. Griffith/Cotter: Tailored-Proteolysis Approach
The Griffith/Cotter method is based on a dual enzyme digestion to reduce the size
of the isopeptide tag, and taking advantage of data-dependent MS/MS acquisition mode
to provide an accurate and fast analysis.108 Data dependent MS/MS is an automated
48
procedure that consists in fragmenting the next three peaks with the highest intensity, and
collecting the MS/MS spectra. Once this step is done, the fragmented parent ions are
excluded from the list and the operation is repeated on the three most intense ions, and so
on, until all the ions have been fragmented. The technique relies on the use of
chymotrypsin, an enzyme that cleaves proteins C-terminal to large hydrophobic amino
acid residues (tyrosine, tryptophan, phenylalanine, and to a lesser extent leucine). In the
case of SUMO-modified proteins, chymotrypsin is of high interest as it yields a 6 amino
acid tag for all human SUMO isoforms (italicized in Fig. 3-4). Because the tag is much
smaller, the modified peptides are more likely to produce quality MS and MS/MS data.
Moreover, mutagenesis is unnecessary to produce a short tag with this approach,
endogenous proteins can be used and the relevance of the analysis is not questionable.
SUMO-2_HUMAN
SUMO-3_HUMAN
SUMO-4_HUMAN
SUMO-1_HUMAN
QIRFRFDGQP
QIRFRFDGQP
QIRFRFGGQP
SLRFLFEGQR
INETDTPAQL
INETDTPAQL
ISGTDKPAQL
IADNHTPKEL
EMEDEDTIDV
EMEDEDTIDV
EMEDEDTIDV
GMEEEDVIEV
FQQQTGG
FQQQTGG
FQQPTGG
YQEQTGG
Figure 3-4: Sequence alignment of the human SUMO C-terminal regions. The tag
obtained by digestion with trypsin and chymotrypsin are shown in bold and italic,
respectively.
The workflow of the Griffith/Cotter method is as follows: the sample containing
several proteins is dually digested with chymotrypsin (for 3 or 4 hours), followed by
trypsin (overnight). The resulting peptide mixture was separated by LC, connected to an
ESI-MS instrument. A survey scan was carried out on the eluted peptides, and they were
then fragmented in a data dependent fashion. One of the features of this technique is the
49
ability to extract and monitor given masses for the daughter ions. This is of interest
because when fragmented, the tag on modified peptides produces ions that are specific to
only the SUMO isopeptide tag. Extracted ion chromatograms from fragment ions
resulting from amide bond cleavage within this QQQTGG- tag can be used to screen
LC/MS/MS data for possible SUMO isopeptides. These ions, named b2’, b3’ and b4’, as
shown in Fig. 3-5 are produced by fragmentation within the SUMO isopeptide tag.
Interestingly, it has been reported that N-terminal glutamine residues can undergo an
internal rearrangement leading to the neutral loss of ammonia (Fig. 3-6).109 Because all
SUMO tags feature an N-terminal Q residue, simultaneous screening for b2’− 17 (240.10
m/z), b3’− 17 (368.15 m/z) and b4’− 17 (469.19 m/z) can selectively point to SUMO
substrate candidates. These ions are found in any SUMO-2 or -3 peptides regardless of
the charge or extent of the proteolytic digest. Consequently, when all three specific
diagnostic mass tag ions are present at the same elution time, there is a high likelihood
that the peptide being analyzed is modified by SUMO.
Figure 3-5: Schematic of an isopeptide carrying the 6 amino acid residues SUMO tag
obtained by dual digestion with trypsin and chymotrypsin. The fragment ion series
obtained by CID are labeled using the Roepstorff and Fohlman nomenclature. The prime
refers to fragment produced by the SUMO tag and not the peptide itself.
50
H2N
H2C
H2N
CH
C
O
CH2
CH2
C
- NH3 (17 Da)
O
NH
O
C
N
H
CH2
CH
C
NH
R
O
R
glutamine
pyroglutamate
Figure 3-6: Scheme of the N-terminal glutamine rearrangement into pyroglutamate with
neutral loss of ammonia.
There are many advantages to using this method: first, it allows faster analysis as
the total time needed for the experiment is on the order of minutes. Because most of the
steps are automated, the user-induced error is reduced. The procedure is also applicable
for all human SUMO isoforms with minimal tweaking to account for differences in the
sequences of SUMO-1 (QEQTGG) and SUMO-4 (QQPTGG). LC separation of peptides
though is very time- and solvent-consuming, leading to current interest in the
development of pre-screening methods to complement the established methods.
3.3. Requirements for a New Method
All current methods for SUMO analysis require steps for separation and/or preconcentration of the analytes prior to MS analysis. These techniques rely on a
compromise: the generation of shorter tags through mutagenesis that are more easily
analyzed by MS is at the expense of physiological relevance. Also higher-throughput
experiments are effective for simple mixtures and become increasingly inefficient for
51
more complex samples. New and effective methods for SUMOylation analysis must
address a number of important factors. The first is the ability to carry out the analysis
using common MS platforms. Secondly, the new method must be fast, efficient and
require less time and solvent; and it has to be a routine procedure that can generate
reliable results. Finally, it must have the ability to analyze SUMOylation in endogenous
human proteins, with no mutatagenesis or genetic tag. The work presented in this thesis
attempts to address these goals and investigates a new method based on ion mobility mass
spectrometry for the screening of SUMOylated proteins.
52
Chapter 4: Material and methods
4.1. Materials
All chemicals used in this study were of analytical grade or better. Solutions of 1.5
M Tris HCl pH 8.8, 1 M Tris HCl pH 6.5, Laemmli sample buffer, 10x Tris/Glycine/SDS
running buffer and 30% acrylamide solutions were purchased from Bio-Rad Life Science
(Hercules, CA). Sodium bicarbonate, ammonium bicarbonate (AmBic), trifluoroacetic
acid (TFA), bovine serum albumin, sulfophenyl isothiocyanate (SPITC) power and
proteomics grade trypsin were purchased from Sigma (St. Louis, MO). Sequencing grade
bovine pancreas chymotrypsin was purchased as a salt free lyophilizate from Roche
Applied Science (Indianapolis, IN). Poly-SUMO2 and -3, SUMO protein set, SUMO-1
conjugation kit, SUMO conjugation substrate UBE2K (E2-25K), UBE2I (UbcH9) and
SUMO activating enzyme (SAE1/SAE2) were purchased from Boston Biochem, Inc.
(Cambridge, MA). GST-tagged RanGAP1 and SP100 fragments (human recombinant,
GST-tagged) were obtained from Enzo Life Sciences (Farmingdale, NY). C-terminal
amidated CCK-8 desulfated peptide (DSpp) and its sulfated analog (Spp) were purchased
from Research Plus (Barnegat, NJ). Lyophilized -casein was purchased from USB
(Cleveland, OH). OMIX 10 μL, C18 resin ziptips were purchased from Varian. All
solvents were of HPLC grade.
53
4.2. In vitro SUMOylation of Proteins
In vitro SUMOylation of protein substrates was carried out in 0.6 mL low
retention Eppendorf microcentrifuge tubes using a SUMOyation kit from Boston
Biochem according to manufacturer’s recommended protocol. The amounts added and
final concentrations for each reagent are presented in Table 4.1. Proteins/reagents were
introduced in the same order as they appear in Table 4.1. The conjugation reactions were
incubated at 37 °C for 3 h.
Table 4-1: Protein and reagent amounts and final concentrations used for in vitro SUMO
conjugation reactions.
Reagent
Final concentration
Amount for a 20 L
reaction
E1 enzyme
100 mM
4 L of 0.5 M solution
SUMO-1/-2/-3
50 M
4 L of 250 M solution
Reaction buffer (10X: 500
mM Hepes pH 8, 1000
mM NaCl, 10 mM DTT)a
1X
2 L of 10X solution
UbcH9
5 M
2 L of 50 L solution
Substrate protein
As specified
5.2 g
Distilled water
N/A
Complete to 20 L
Mg-ATP
1 mM
2 L of 10 mM solution
a
DTT = dithiothreitol
4.2. SDS-PAGE Gels
To verify the extent of the modification after in vitro SUMOylation, one fourth of
the volume of each reaction was run on a 15% and 5% acrylamide/bis acrylamide SDS54
PAGE separating and stacking gels, respectively. The gels were prepared as reported in
literature.110 Table 4.2 summarized the volumes of the various reagents used in
preparation of the gels. The 10% ammonium persulfate solutions were freshly made prior
to use.
Table 4.2: Chemicals and amounts used for the preparation of 15% acrylamide SDSPAGE gel.
15% resolving gel (10 mL)
5% stacking gel (2 mL)
Distilled water
2.3 mL
1.4 mL
30% acrylamide mix
5.0 mL
0.33 mL
1.5 M Tris (pH 8.8)
2.5 mL
-
1 M Tris (pH 6.5)
-
0.25 mL
10% SDS
10% ammonium
persulfate
TEMED
0.1 mL
0.02 mL
0.1 mL
0.02 mL
0.004 mL
0.002 mL
In vitro SUMOylation sample mixtures were mixed in a 1:1 ratio with Laemmli
sample buffer and boiled for five minutes before introduction into the wells. The gels
were run at 200 V for 80 min using 1x Tris/glycine/SDS buffer. The masses of the
proteins were estimated in comparison to the migration of a standard protein ladder
(Precision Plus Protein Dual Xtra Standard), ranging from 2 kDa to 250 kDa.
4.3. Enzymatic Digestion
A sample containing 5 L of a 2.2 g/L poly-SUMO-2 or -3 solution (provided
in 50 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) pH 8.0, 100 mM
55
NaCl, 1 mM DTT), or the products of in vitro SUMOylation were evaporated to dryness
by Speed-Vac, and reconstituted in 20 L of 50 mM AmBic pH 8.5 for digestion.
Sequencing grade chymotrypsin (1 μL of 1 mg/mL solution in 1 mM HCl) was added in a
1:50 enzyme/substrate ratio, and the solution was incubated for 4 h at room temperature
(25 °C). Proteomics grade trypsin was then added (1:25, 2 μL of 1 mg/mL solution in 1
mM HCl) and incubated overnight (approximately 15 h) at 37 °C. Digestion was
quenched by the addition of formic acid to a concentration of 0.1% v/v. Aliquots (10 μL)
of the digest sample were desalted using 10 μL C18 ziptip following the manufacturer’s
recommended protocol (as described in section 4.4.) and subsequently evaporated to
dryness by Speed-Vac for MS analysis.
4.4. Zip Tip Procedures
4.4.1. Manufacturer Protocol
SUMOylated, digested substrates and sulfated/desulfated peptides were desalted
using 10 μL C18 ziptip following the manufacturer’s recommended protocol and
evaporated to dryness by the Speed-vac. In short, five solutions were prepared: wetting
solution (5% acetonitrile (ACN) in water), equilibration and wash solutions (0.1% TFA in
water), elution solution 1 (0.1% TFA in water containing 50% ACN) and elution solution
2 (0.1%TFA in water containing 50% ACN). Using 10 L Eppendorf pipette and the
Ziptip media was wet with wetting solution. Equilibration was then achieved by
aspiration of the equilibration solution. Peptides were bound with 7 to 10 slow aspirate
and dispense cycles of the sample solution. The tip was rinsed with the wash solution,
which was discarded in waste. Peptides were recovered with 15 aspirate and dispense
56
cycles with 20 L of elution solution 1, followed by the same step with elution solution 2.
The two solutions are finally combined and sample is immediately dried with Speed-Vac.
4.4.2. Preparation of Oligonucleotides Procedure
N-terminal sufonated peptides were desalted using 10 μL C18 ziptip using a
modified procedure optimized for better retention of oligonucleotides and negatively
charged analytes. Five solutions were prepared for the procedure: wetting (50%
acetonitrile in water), equilibration (0.1 M triethylammonium acetate (TEAA), pH 7.0),
wash solution #1 (0.1 M TEAA, pH 7.0), wash solution #2 (water) and elution buffer
(50% ACN in water). The sample was dissolved in 10 L of the equilibration solution
prior to desalting. The Ziptip was wet using the wetting solution, and then equilibrated
with 10 L of equilibration solution 3 times. Peptides were bound with 5 to 10 aspirate
and dispense cycles of the sample solution. The tip was rinsed 3 times with 10 L of both
wash #1 and #2 solutions. Finally, peptides were recovered by at least 3 cycles of
aspiration and dispensing into 10 L of the elution buffer. Desalted samples were
immediately dried using Speed-Vac.
4.5. Mass Spectrometry
4.5.1. Nano-ESI-TOF/IM MS
Desalted digested samples were reconstituted in 50 μL of a solution containing
0.05% formic acid and 50% ACN for MS analysis. All mass spectrometry data were
collected on a Synapt HDMS quadrupole time-of-flight ion mobility mass spectrometer
equipped with a nanospray source (Waters Corp.). The following instrumental parameters
were used for all experiments: capillary, 2.5-2.8 kV; sampling cone, 40.0 V; extraction
57
cone, 4.0 V; cone gas flow rate, 0 L/h; trap collision energy (CE), 6.0 V; transfer CE, 4.0
V; trap gas flow, 1.5 mL/min; IMS gas flow reate, 0 mL/min. For ion mobility
experiments, the parameters were kept identical with the exception of IMS gas flow (2027 mL/min). For MS/MS experiments the trap CE was increased to 15.0-35.0 V. Mass
spectra were calibrated externally in the positive ionization mode in the range 500 ≤ m/z
≤ 4000 using a solution of sodium cesium iodide, and processed using Masslynx 4.1
software (Waters). All mass spectra were averages of approximately 300 scans and
presented unprocessed (unsmoothed and without background subtraction). Ion mobility
data were processed using the Driftscope 2.0 software.
4.5.3. MALDI MS
The progress of the BSA digestion was monitored using the UltrafleXtreme
MALDI-TOF/TOF MS equipped with a smartbeam II laser (Bruker Daltonics). A
supersaturated solution of -cyano-4-hydroxycinnamic acid (CHCA) in 0.05% TFA
containing 50% ACN was prepared. The sample was spotted onto a stainless steel plate
using the sandwich method where 1 L of matrix solution was spotted and dried,
followed by 1 L of sample solution, and then another 1 L of matrix solution. Data were
collected in the positive ionization and reflectron modes. External calibration was done in
the range 400-4000 m/z using angiotensin II, 1056.5418 m/z; angiotensin I, 1296.6848
m/z; Substance P, 1347.7354 m/z; bombesin, 1619.8223 m/z; ACTH clip 1-17, 2093.0862
m/z ACTH clip 18-39, 2465.1983 m/z; and somatostatin 28, 3147.4710 m/z (Peptide
calibration standard #206195, Bruker Daltonics). Approximately 3000-8000 shots were
acquired per MS spectrum using 1000 Hz acquisition speed. FlexAnalysis 3.3 software
58
(Bruker Daltonics) was used for data processing. The peaks obtained were used to carry
out a peptide mass fingerprinting with the MASCOT software.
4.6. In silico Digests
The mass lists of the expected peptides from the tryspin/chymotrypsin digested
substrates were generated in silico from the protein sequence using the “MS-digest”
function of Protein Prospector, a free utility created by the University of California, San
Francisco (prospector.ucsf.edu/prospector/mshome.htm). The calculated masses of
peptides were compared against the experimental masses in order to match sequences.
4.7. Spiking of BSA Digests
0.001 g of bovine serum albumin (BSA) were digested in 21 L of 50 mM AmBic
pH 8.5 with 19 L of 1 mg/mL trypsin solution; total volume 40 L and final BSA
concentration 500 M. The solution was incubated at 37 °C for two days. Working
samples were prepared by spiking the appropriate volume of a 87 L solution of Spp to
achieve a 5 M solution of BSA digest containing 1:1 molar ratio of Spp.
59
Chapter 5: Pre-Screening Method for SUMOylated
Proteins
5.1. Proof-of-Concept for the Screening Method
5.1.1. Overview
As stated previously, one of the project goals was to remedy the lack of rapid and
efficient methods to confidently analyze with confidence SUMOylated protein. Due to its
separation properties, IMMS was chosen to distinguish between the modified and
unmodified peptides, allowing removing of any purification or pre-concentration step
prior to the MS analysis. A facile method was developed for screening simple protein
digests for possible modification by SUMO. With the use of poly-SUMO2 as a model
(sequence shown in Fig. 5-1), the method includes digestion with trypsin/chymotrypsin,
followed by ion mobility mass spectrometry analysis.111 Using a two-enzyme system
typically leads to small peptides. The presence of the QQQTGG-tag results in a
significant increase in mass (+ 618 Da) and size relative to the unmodified peptide. We
exploit denaturing solution conditions to promote higher charge states; and IMMS to
separate charge states in order to screen/identify possible SUMO isopeptides, which are
confirmed by MS/MS. This method is very simple and much faster than LC-MS/MS
approaches.
60
1 MADEKPKEGV KTENNDHINL KVAGQDGSVV QFKIKRHTPL SKLMKAYCER
51 QGLSVRQIRF RFDGQPINET DTPAQLEMED EDTIDVFQQQ TGG
Figure 5-1: Sequence of human SUMO-2. Sequence covered by MS is indicated by
underlined. In bold red is shown the SUMO-modified lysine residue.
5.1.2. ESI-MS Analysis of the Poly-SUMO-2 Digest
SUMO-2 possesses the interesting feature of being able not only to modify its
substrate, but also to be modified by itself at K11, which is found within a SUMOylation
consensus sequence. This results in the formation of polymeric chains linked by
isopeptide bonds. A solution of poly-SUMO-2 containing chains of various lengths was
used as a model system. The mass spectrum of the trypsin/chymotrypsin digest of polySUMO-2 is provided in Figure 5-2. The vast majority of these peptides and also the most
abundant peptides could be matched to theoretical masses for the trypsin/chymotrypsin
digest calculated from the sequence of SUMO-2. The sequence covered by the identified
peptides is shown as underlined in Fig. 5-1. It is interesting to note that peptides yielding
from cleavage after the substrate lysine are obtained, although it is known that modified
peptides exhibit a missed cleavage site at the modified lysine. It is actually due to the fact
that the SUMO molecule at the end of the polymeric chain is not modified. Thus, the
lysine C-terminal amide bond can be cleaved by trypsin and peptides starting at T12 can
be observed. The peptides detected range in charge state from +1 to +3. Under the
solution conditions used (50% acetonitrile, 0.05% formic acid), relative to neutral
aqueous solutions, it is expected that peptides will acquire a larger number of charges,
depending on their size and in some part sequence. All of the larger peptides detected had
61
a charge of 3+. As expected for a two-enzyme digest with trypsin and chymotrypsin, the
resultant peptides are relatively smaller in mass than expected with a one-enzyme digest.
A list of peptides detected and the sequence coverage are provided in Table 5.1. The
identity of these peptides was confirmed by MS/MS and the data is presented in
Appendix A-1 to A-7. It was observed that some peaks do not match the expected masses
for the linear SUMO-2 peptides, such as the ones at 1033.04 and 1041.55 m/z.
Figure 5-2: Mass spectrum of the trypsin/chymotrypsin digest of poly-SUMO-2 showing
some of the linear peptides identified. The numbers of the peptides refer to the number
assigned to each peptide in Table 5.1 shown below.
62
Table 5.1: List of the major linear peptides detected from the poly-SUMO-2 dual digest.
These peptides are indicated on the mass spectrum in Figure 5-2. m/z: experimentally
measured m/z value of peptide ion; z: charge of peptide ion; Mcalculated: neutral mass of
peptide calculated from the monoisotopic experimentally measured m/z value; Mtheoretical:
neutral mass of peptide calculated from sequence; ΔM: monoisotopic mass error of
measurement.
#
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
Sequence
position
12-20
22-32
12-21
21-32
22-33
77-87
63-76
62-76
61-76
77-87
22-32
44-53
77-87
63-87
62-87
61-87
m/z
z
Mcalculated
Mtheoretical
M
535.2824
553.8039
599.3212
+2
+2
+2
1068.5648
1105.6078
1196.6424
1068.4909
1105.5477
1196.5858
0.0739
0.0601
0.0566
617.8597
+2
1233.7194
1233.6426
0.0768
671.8137
749.8804
823.4124
901.4847
941.4407
1106.6132
1198.6930
1342.6472
1411.6888
1485.2484
1563.3177
+2
+2
+2
+2
+3
+1
+1
+1
+2
+2
+2
1341.6274
1497.7608
1644.8248
1800.9694
2821.3222
1105.6132
1197.6930
1341.6472
2821.3776
2968.4968
3124.6354
1341.5355
1497.7020
1644.7704
1800.8715
2821.2197
1105.5477
1197.5707
1341.5355
2821.2197
2968.2881
3124.3892
0.0919
0.0588
0.0544
0.0979
0.1025
0.0655
0.1223
0.1117
0.1579
0.2087
0.2462
5.1.3. Ion Mobility Analysis of the Poly-SUMO-2 Digest
A very practical advantage of IMMS is the ability to separate multiply charged
ions generated from electrospray ionization into their respective charge states.50 Figure 53 shows an ion mobilogram (or mass-mobility plot) for the mass spectrum provided in
Figure 5-2. The data is representative of previously reported mass-mobility plots for
protein digests where the majority of ion signals exhibit a high correlation to diagonal
lines
112
. As indicated, the diagonal lines correspond to the various charge states of
peptide ions. The lower trend line passes through peaks for peptides with only one charge.
63
At the resolution of our IMMS instrument, it is possible to separate the +2 and +3 charge
states. Negligible amounts of any higher charge states were observed. Only one linear
peptide (no. 9 in Table 5.1) was detected with the +3 charge state.
Figure 5-3: Ion mobilogram/mass-mobility plot (drift time versus m/z diagram) of the
trypsin/chymotrypsin digest of human poly-SUMO-2. The diagonal trend lines
representative of the various charge states of ions are indicated.
64
Figure 5-4: Drift time total ion chromatogram (black solid trace) of peptides detected
from the trypsin/chymotrypsin proteolysis of poly-SUMO2. Extracted drift time ion
chromatogram of only the larger more highly charged (z ≤ +3) ions (dashed trace).
The utility of IMMS can be seen in Figure 5-4. The drift time total ion
chromatogram (solid black trace) shows the distribution of drift times of all ions detected
in the poly-SUMO2 digest. The gray dashed trace represents the extracted ion
chromatogram of only the larger ions with charge state +3. This grouping of ions, as
expected, has a much narrower drift time distribution. On the basis of the intensity of the
maximum at approximately 2 ms, relative to total ion chromatogram, the +3 ions
represents only a very small percentage all ions in the mass spectrum. Despite the lower
abundance (<35% of the intensity of the base peak), the extracted mass spectrum
representing these +3 charge state ions (Figure 5-5) shows a signal-to-noise ratio
sufficient for definitive determination of the monoisotopic masses of these peptides
(Table 5.2). Using MS/MS, the identities of these peptides were confirmed (Fig. 5-7 and
65
Appendix B-1). SUMO-2 isopeptides are labeled in Figure 5-5. Comparison of the
experimental data with a computer simulation of the isotopic distribution for the
isopeptides increased the confidence of the assignment (Fig. 5-6 and Appendix B-2 to B4). As expected, these peptides correspond to a peptide containing lysine-11 of SUMO-2,
which lies within the consensus sequence, modified by the QQQTGG- isopeptide tag.
Substrate peptides all begin at a trypsin cleavage site, while termination from both trypsin
and chymotrypsin cleavage was observed. The isopeptides terminating from
chymotrypsin cleavage were in much higher abundance than the corresponding peptide
that terminated with a trypsin cleavage site. The sensitivity of the method was high
enough to obtain the monoisotopic peak and several isotope peaks for all the isopeptides,
allowing for unambiguous mass determination. This also explains the peak clusters
observed for each isopeptide.
66
Figure 5-5: Mass spectrum reconstructed from the gray dashed trace in Figure 5-4
showing only the peptides with z ≥ +3. Confirmed SUMO-2 isopeptides are indicated.
Figure 5-6: Comparison of the experimental (blue solid trace) and simulated (red dashed
trace) isotopic distributions for isopeptide 2 in Table 5.2
67
Table 5.2: List of the SUMO-2 isopeptides detected using ion mobility mass
spectrometry. These peptides are indicated on the mass spectrum in Figure 5-5.
#
1
2
3
4
m/z
689.0397
694.6915
731.7465
737.4106
z
+3
+3
+3
+3
Mcalculated
2064.0957
2081.0511
2192.2240
2209.2084
Mtheoretical
2063.9946
2080.9773
2192.0696
2209.0723
M
0.1011
0.0738
0.1544
0.1361
For all species, both [M + 3H+]+3 and [M + 3H+ − 17]+3 ions were observed. This
17 Da loss has been previously shown to occur in peptides containing N-terminal
glutamine residue.109 Under acidic solution conditions, peptides with N-terminal Gln can
undergo internal nucleophilic attack with the expulsion of ammonia, resulting in Nterminal pyroglutamic acid (as shown previously in Fig. 3-6). The b’ − 17 fragment ions
were also shown to be highly abundant in the tandem mass spectra of the N-terminal Qcontaining peptides (Fig. 5-7). Only isopeptide no. 2 of Table 5.2 was also detected with a
+2 charge state. Once potential candidates have been identified, these are confirmed by
tandem mass spectrometry and the presence of diagnostic fragment mass tags for b2’ − 17,
b3’ − 17, and b4’ − 17, which originate from the isopeptide tag.108
68
Figure 5-7: MS/MS spectrum of the isopeptide 731.74 m/z (+3), 3 in Table 5.2. The b’n
fragments originating from the SUMO tag were clearly identified.
5.1.4. Other SUMO Isoforms
Because all SUMO isoforms possess a chymotrypsin cleavage site 6 amino acids
from the C-terminal, the method can be applied to any human SUMO. Following the
same experimental procedure as for SUMO-2, a poly-SUMO-3 sample was analyzed. The
experiment resulted in the detection of 4 isopeptides, both [M + 3H+]+3 and [M + 3H+ −
17]+3 as shown on Fig. 5-8. The satellite peaks observed (above 700 m/z) correspond to
sodium adducts, and are present due to inefficient Ziptip desalting of the sample. Because
SUMO-1 and -4 have not been reported to form polymeric chains in vivo, the method
could not be tested against the polymers of those two isoforms. However, due to the high
69
similarities of the tag obtained from SUMO-1 and -4, it is expected that this method will
be suitable for all human isoforms.
Figure 5-8: Mass spectrum reconstructed showing only the peptides with z ≥ +3 for the
dually digest poly-SUMO-3 sample. Confirmed SUMO-3 isopeptides are indicated.
5.2. Applications of the Method
5.2.1. In vitro SUMOylation: Sp100 and RanGAP1
In an attempt to increase the complexity of the sample analyzed, and get closer to
the goal of studying a complete cell lysate, in vitro SUMOylated proteins were
investigated. The reaction was conducted on two reported SUMO-1 substrates, Sp100 and
RanGAP1 fragments, whose sequences are presented in Fig. 5-9.
70
1 INLNDNTFTE KGAVAMAETL KTLRQVEVIN FGDCLVRSKG
41 AVAIADAIRG GLPKLKELNL SFCEIKRDAA LAVAEAMADK
81 AELEKLDLNG NTLGEEGCEQ LQEVLEGFNM AKVLASLSDD
1 KAEPTESCEQ IAVQVNNGDA GREMPCPLPC DEESPEAELH
41 NHGIQINSCS VRLVDIKKEK PFSNSKVECQ AQARTHHNQA
81 SDIIVISSED SEGSTDVDEP LEVFISAPRS EPVINNDNP
Figure 5-9: Sequences of the commercial RanGAP1 (top) and Sp100 (bottom) fragments.
In red are shown the reported SUMO-modified lysine residues found in the consensus
sequence.
Due to the presence of not only one protein and two enzymes (as for the
experiments described above), the sample contained SUMO-1, the substrate protein, E1
and UbCh9 enzymes required for the reaction, the small molecule Mg-ATP and the two
proteases (trypsin and chymotrypsin). For the reaction to yield a sufficient amount of
modified protein, SUMO-1 had to be introduced in large excess compared to the substrate
(1:10 ratio substrate:SUMO-1). Because of this, and the many species are present in the
sample, the percentage of modified peptides compared to the total number of peptides in
the sample is significantly decreased. Moreover, in vitro SUMOylation produces less
modified protein compared to auto-SUMOylation of SUMO-2 and -3. Consequently,
using in vitro SUMOylated proteins is a good way to test for the sensitivity and
robustness of the method. The gel obtained after the reaction for Sp100 and RanGAP1 is
presented below in Figure 5-8. The first striking observation is that the commercial
fragments, analyzed as received, displayed multiple bands on the gel. This is especially
notable for the Sp100 sample that seems to contain multiple species. There is no evidence
on this gel for conjugation of SUMO-1 with either Sp100 or RanGAP1. The bands
obtained for the complete reaction mixture and the controls are the same, showing that no
new products are formed by the in vitro SUMOylation reaction.
71
Figure 5-10: SDS-PAGE of in vitro SUMOylation of Sp100 and RanGAP1 fragments in
presence or absence of SUMO-1 and ATP.
MS experiments (MS scan and IMMS) on the dual enzyme digested and desalted
samples did not lead to any definitive results. About one half of the peaks obtained on the
TOF spectra were matched with SUMO-1 peptides; however none of the masses matched
the expected sequence for the Sp100 and RanGAP1 fragments, or any of the enzymes that
were present in the solution (Appendix C-1 and C-3). Trypsin digestion of the
commercial substrates followed by MALDI analysis (Appendix C-2 and C-4) and peptide
72
mass fingerprinting revealed that the substrate sequence was not the one indicated by the
manufacturer. GST-tagged Sp100 fragment digested with trypsin led to a MASCOT hit
for GST-tag only. None of the other detected peaks matched the theoretical m/z list
generated in silico, confirming that the wrong proteins were provided by the
manufacturer. Moreover, the same experiment was carried out on the GST-tagged
RanGAP1 fragment, which did not score any hit with the MASCOT search. Manual
matching of the obtained peaks revealed that once again, the sequence of the fragment is
not likely to be what was indicated by the manufacturer. Because of this issue,
experiments to apply the method were delayed.
73
Chapter 6: Analysis of Tyrosine O-Sulfation by IMMS
6.1. Physiological Functions of Small PTMs
6.1.1. Sulfation
Sulfation refers to chemical or enzymatic modification by covalent addition of a
sulfate moiety (SO42-). Biologically, this reaction is an irreversible post-translational
modification in vivo113 that targets the hydroxyl group on tyrosine residues in proteins
(termed tyrosine O-sulfation) leading to the modified amino acid, sulfotyrosine. The
identification of sulfated proteins coupled to the mapping of their modification sites led to
the conclusion that there is no consensus sequence for tyrosine O-sulfation.114 Although
some have described consensus features (tyrosine residues exposed on the surface of the
protein and typically surrounded by acidic residues), many protein sulfation sites do no
fulfill those features.115 The first occurrence of protein sulfation was discovered in 1954
in a peptide derived from bovine fibrinogen.116 The first human proteins found to carry a
sulfotyrosine (7 of them) were reported in 1985 by Liu et al.117 Sulfation is catalyzed in
vivo by the tyrosylprotein sulfotransferase (TPST) enzymes in the Golgi apparatus.118 It
was later discovered that two variants of this enzyme exist, TPST-1 and -2, which display
high homology of both sequence and structure.119
74
More than 275 sulfated proteins have been discovered and identified, mostly
secreted proteins and trans-membrane spanning proteins. However, the exact biological
functions of tyrosine O-sulfation remain unclear. It is thought that this PTM is a key
regulator for protein-protein interactions, it even seems that some interactions are made
through recognition of the sulfate group itself.120 Some of the known mechanisms in
which sulfation plays a role include the modification of the chemokine receptor,
involvement in the entry events of human immunodeficiency virus 1 (HIV-1) in target
cells; leukocyte adhesion and inflammatory response; homeostatis (modification of
coagulation factors forming a stable complex); and modification of neurologically
bioactive peptides.121 The biological function and the significance of some of these
modifications are currently still unknown.
Various methods for analysis of sulfated proteins have been reported
previously.122 Historically, metabolic labeling with 35S-sulfate was used for the screening
of sulfated substrates123 and is still currently used as a back-up method for unambiguous
identification of sulfotyrosine residues. The main disadvantage is the non-specificity of
this labeling technique; it does not distinguish between actual tyrosine sulfation, and
sulfation of protein linked carbohydrates. Separation of sulfated peptides is typically
carried out by RP-HPLC, the modified species elute more rapidly than their unsulfated
counterparts.122 Some spectroscopic methods are used for characterization of
sulfotyrosine peptides; UV spectroscopy is especially efficient for quantification as the
presence of a sulfate group dramatically changes the absorbance features of the aromatic
ring: shift of the absorbance maximum from 275 nm to 260.5 nm in 0.01 M HCl or from
293 nm to 263 nm in 0.01 M NaOH.116 Fourier transform infrared spectroscopy is also a
75
common technique for the analysis of sulfated proteins and peptides. Indeed, the
symmetric SO3 stretching vibrations give rise to an intense band at 1050 cm-1;
supplementary bands corresponding to the asymmetric vibrations can also be observed at
1230 cm-1 and 1270 cm-1.124 Despite the wide range of available analytical instruments for
the characterization of sulfation, the current technique of choice is MS. MALDI and ESI
sources have made it possible to analyze labile modifications, and MS/MS data allow for
accurate mapping of the modification sites. This is usually achieved by methods including
enzymatic digestion and fragmentation methods such as CID. However, this workflow is
time-consuming and requires relatively large quantities of protein. With the introduction
of ECD, that preferentially breaks peptide backbones, better sequence coverage and better
retention of PTMs was obtained. One of the main difficulties in MS analysis of sulfated
proteins is that the mass added by a sulfate and a phosphate group are almost identical.
Methods to distinguish between these two PTMs are presented in more detail in Section
6.1.3.
6.1.2. Phosphorylation
Phosphorylation is the covalent addition of a phosphate group (PO43-) to a
substrate. Unlike sulfation, phosphorylation is reversible and can modify several amino
acid residues. Serine, tyrosine, and threonine display the most abundant phosphorylation,
but low stability modifications of aspartic acid, glutamic acid and histidine are also
observed.72 The phosphorylation of proteins is regulated by a balance between the activity
of kinases (enzymes that catalyze phosphorylation) and phosphatases (enzymes that
remove the phosphate group from modified proteins). The human genome encodes for
500 kinases and 100 phosphatases, revealing the importance of this modification. It is
76
estimated that approximately 30% to 50% of proteins are phosphorylated at some point in
their lifespans.125 However, at any given point in time, only 1% of the proteins in the cell
are phosphorylated.126 Phosphorylation is involved in a large number of mechanisms, and
is widely used in the cell for regulation of protein stability, localization, function and
activity.127 Moreover, this PTM controls many cellular processes such as cell division,
enzymatic
activity,
signal
transduction,
and
metabolic
pathway regulation.128
Deregulation of the phosphorylation machinery is involved in several diseases, including
cancer.129
6.1.3. Sulfation, Phosphorylation: Similarities and Differences
A summary of some characteristics of phosphorylation and sulfation as PTMs is
presented in Table 6.1. Sulfation and phosphorylation are often thought to be homologous
modifications, especially from a MS perspective, as they both add 80 Da to the mass of
their substrates and are highly acidic. Although sulfur and phosphorus have different
atomic masses (32.065 and 30.974 respectively), a phosphate group carries 3 negative
charges while a sulfate group only has 2. Thus, phosphorylation requires ion pairing with
one proton to result in a single negative charge, which is not the case for sulfation.
Because of this additional proton, the mass of the modification is increased by 1.008,
making phosphorylation and sulfation isobaric. However, the nature of the modification,
the physiological functions and the behavior in the MS instruments are different for
sulfation and phosphorylation. Phosphopeptides can undergo several losses during CID,
which are the PO3- ion (− 80 Da), neutral HPO3 (− 80 Da) and H3PO4 (− 98 Da) or H3PO4
+ H2O (− 116 Da).130 Loss of H3PO4 occurs readily and typically generates a high
intensity peak; it has to be noted that the neutral loss of phosphoric acid occurs for pS and
77
pT only. CID of sulfopeptides only leads to the loss of the sulfate ion SO3- (− 80 Da).131
Sulfation and phosphorylation groups actually have different monoisotopic masses
(respectively 79.9568 Da and 79.9663 Da) and can be distinguished without ambiguity
when the resolution of the instrument is high enough to attain mass accuracies of 5 ppm
or better. This approach is usually not applicable as the FT-ICR instruments that provide
enough resolution are not available in many MS facilities. Another alternative is to use a
combination of positive and negative ionization: peptides usually lose their sulfo moiety
in positive mode while phosphopeptide intact ions can be obtained in both ionization
modes. Furthermore, in negative ionization mode phosphorylation produces a specific
negative ion at 79 m/z whereas sulfation forms an equivalent ion at 80 m/z.
78
Table 6.1: Comparison of some biological and biochemical features of phosyphorylation
and sulfation as PTMs; together with their properties during MS analysis.
Biology
Phosphorylation
Sulfation
Intracellular
Extracellular
Reversible
Irreversible
Activation, inactivation,
modulation of protein
interactions
Modulation of
protein interactions
Location
Reversibility
Function
Biochemistry
MS
Chemical
stability
pY is stable
pS/pT are alkaline labile
sY is acid labile
pE/pD/pH are acid labile
Removal
Phosphatases
Arysulfatases
Property
Acidic (2 −)
Acidic (1 −)
Monoisotopic mass
change
+ 79.9663 Da
+ 79.9568 Da
pS/pT: good
pY: stable
sY: easily lost under
standard conditions
PO3- (− 79.9663 Da)
SO3- (− 79.9568 Da)
Stability during
ionization
Characteristic
fragment loss
6.2. Project Goal
As shown in the previous sections, sulfation and phosphorylation are involved in
many biological mechanisms, and complete understanding of their functions requires
identification of the modified proteins. The goal of this project was to apply IMMS to the
analysis of sulfated sites in proteins, using approaches based on currently available
methods for phosphorylation analysis.
79
The addition of a sulfate or phosphate group to a peptide triggers a change in its
formal charge and general structure. These changes can be exploited by using ion
mobility MS, as it provides separation of ions not only based on their m/z but also formal
charge, size and shape. Due to the increase of the negative charge and change of structure
of the modified peptides, it was hypothesized that the drift time of these species would be
altered. Consequently, the band for these ions in the correlation lines on the ion
mobilogram plot would be slightly shifted, making the screening for modified peptides
fast and straightforward. It was extensively demonstrated in the past that phosphorylated
peptides exhibit a more compact structure that shifts their drift times to lower values.112,
132
Because the behavior of phosphopeptides in IMMS is so well characterized, they were
chosen as model system for understanding the effects of each instrumental parameter on
the shift of the modified species.
Bovine -casein protein was chosen as a model for phosphorylation, and used in
IMMS experiments to optimize the experimental conditions. This established approach
was then extended to sulfation with the use of an eight-residue peptide in combination
with its sulfated analog.
6.3. Optimization of the IMMS Experiments: Phosphorylation
6.3.1. Model System
Casein is the major protein of bovine milk and is well known for being a
phosphorylation substrate. Casein is found as a heterogeneous mixture of three isoforms:
-, - and -casein present at 75%, 22% and 3% respectively. Alpha casein is constituted
of 214 amino acid residues (24,529 Da) and can be phosphorylated at multiple serine
80
positions: 56, 61, 63, 79, 81, 82, 83, 90, 130.133 It was used as model system to study
phosphorylation as modified protein is available commercially. This protein forms
hydrophobic micelles and is not very soluble in water and aqueous solutions.
6.3.2. Results
A 4.8 mg sample of multiply phosphorylated -casein lyophilized powder was
dissolved in 400 L of ammonium bicarbonate. Because of the poor solubility of the
protein, the pH was increased to about 9 and 40 L of 50 mM magnesium acetate was
added. The protein eventually dissolved and the solution had a final volume of 696.4 L
with a pH of about 9. From the stock solution, 25 L was digested overnight with trypsin
and after a 1:33 dilution in denaturing solvent was analyzed by ESI-TOF MS (Fig. 6-1).
The spectrum is presented after processing by the MaxEnt3 function, which converted all
the peaks to the +1 charge state. Several unmodified and modified peptides were
identified; interestingly, two pairs of identical peptides differing by only one
phosphorylation were found. The extensive list of -casein peptides identified is
presented in the Appendix D-1. Tandem MS on the precursor peak 1660.7 m/z confirmed
the sequence of a phosphorylated peptide 121-134, as shown in Figure 6-2.
81
Figure 6-1: ESI-TOF MS spectrum of an -casein trypsin digest in 50 mM AmAc pH
7.0. The displayed spectrum was processed using the MaxEnt3 algorithm.
82
Figure 6-2: MS/MS spectrum of the precursor ion 1660.70 m/z corresponding to the
singly phosphorylated peptide VPQLEIVPNsAEER. pS represents a phosphorylated
serine residue.
As the TOF and MS/MS data confirmed the presence of phosphorylated peptides,
IMMS experiments were carried out on the digest mixture (Fig. 6-3). The mass spectra of
bands that displayed a shift in their mobility from to the charge correlation lines were
obtained for peptide identification. Two phosphorylated peptides were identified (106119 and 43-58), the second one carrying two modifications. The identity of the peptides
was confirmed by MS/MS. Complete sequence information was not obtained for most of
these peptides.
83
Figure 6-3: Mass-mobility plot of the -casein digest in AmAc pH 9. In bands indicated
by the arrows correspond to phosphorylated peptides and display a shift from the
correlation lines.
The two confirmed phosphorylated peptides displayed a shift from the 2+
correlation line, as shown in Fig. 6-3. Interestingly, the doubly modified peptide shows a
much more marked shift in drift time compared to the mono-phosphorylated peptide. As
stated previously, there is a structural difference between the phosphorylated and nonphosphorylated peptides. This can be attributed to the intra-molecular interactions
between the partial negative charge carried by the modification and basic sites (arginine,
lysine, histidine side chains and protonated N-terminus) that are positively charged.132
The presence of two modifications on a single peptide is likely to increase the strength of
these electrostatic interactions thereby making the structure more compact. As the
experiments with phosphopeptides produced the expected results, it was concluded that
84
the instrument was suitably optimized and the approach was then extended to the sulfated
peptides.
6.4. Screening for Sulfated Peptides
6.4.1. Model System
To test the performance of IMMS for the screening and identification of sulfated
peptides, the model system chosen consisted of a fragment of cholecystokinin, a protein
secreted in the small intestine and known sulfoprotein. The sequence of the peptide is as
follows: DYMGWMDF-NH2 (C-terminus is amidated) and two samples were purchased,
differing only by a sulfate group on the tyrosine. The non-modified peptide is referred to
as desulfated peptide (DSpp) in contrast to the sulfated peptide (Spp). Their monoisotopic
masses were 1063.4012 and 1142.3580 Da, respectively.
6.4.2. Analysis of Spp and DSpp
An equimolar mixture of Spp and DSpp was prepared, subjected to N-terminal
sulfonation and analyzed in denaturing solution by ESI-TOF MS after desalting. The
spectrum obtained showed not only salt contamination, but also extremely small intensity
signal for the sulfated peptide. It was hypothesized that, the peptides (especially Spp) had
a quite high negative charge and did not bind efficiently to the reverse phase C 18 column.
Another approach was attempted, using a zip tip protocol optimized for oligonucleotides,
which are also negatively charged species.
IMMS analysis of the Spp solution only (Fig. 6-4) showed very interesting
features. The correlation line for the +2 charge state was obtained from the BSA
experiment that was done under the same conditions (see Fig. 6-7). Firstly, the desulfated
85
peptide is clearly present in the solution although the sample contained only Spp. The
identity of the peptide was confirmed by MS/MS (Fig. 6-5). This demonstrated the
lability of this modification in the mass spectrometer. MS/MS data could not be obtained
for Spp since any increase of the collision energy (required to fragment the analyte) led to
the loss of the modification. It can also be observed that the non-modified peptide is on
the charge correlation line, meaning that the drift time is consistent with its m/z value.
However, the modified Spp peptide band was to the left of the correlation line, indicating
that its drift time was shifted to smaller values. This observation is consistent with the
behavior of phosphorylated species in IMMS: it is very likely that the negative charge of
the sulfate group also interacts with the positively charged N-terminus, making the
overall structure more compact. The other bands found on the mass mobility plot are
likely to be the same peptides (same m/z) but with different and more compact
conformations (much smaller drift time) that exist in equilibrium.
86
Figure 6-4: IMMS analysis of a solution of Spp in denaturing solution
Figure 6-5: MS/MS spectrum of the 1063.41 m/z (+1) peptide. The obtained sequence
confirmed the identity of DSpp.
87
6.4.3. Spiking of a Bovine Serum Albumin Digest with Spp
As the IMMS analysis of a simple modified peptide showed interesting results,
another experiment was designed to test for sensitivity of IMMS for detection of the
sulfated peptide. Bovine serum albumin was digested with trypsin and mixed in 1:1 ratio
with Spp to a working solution concentration of 5 M. Comparison of the experimental
values with an in silico generated list of BSA peptides (Fig. 6-4 and Appendix E-1)
allowed all the main peaks to be positively identified as BSA tryptic peptides. The Spp or
DSpp peaks were not detected in TOF mode.
Figure 6-6: Positive ESI-TOF MS spectrum of a 5 M BSA digest spiked with a 1:1
molar ratio of Spp.
88
IMMS experiment were carried out on the same sample and although the 1142.36
peak (+1 charge state for Spp) was totally absent, a +2 peak at 571.75 m/z was detected,
only by IMMS and not by TOF-MS. MS/MS could not confirm that the peptide was
carrying a modification since the instrumental parameters required to obtain
fragmentation of the peptide led to its desulfation (i.e. only the desulfated mass was
observed). On the mass-mobility plot, the 571.75 peak showed a shift in drift time to the
left, as indicated on Figure 6-5. Another band for the +2 charge state peptide of Spp at
674.2 m/z displayed a large shift in drift time compared to the correlation line; the
calculated mass matched a methionine-oxidized peptide from the BSA digest. The exact
explanation for this behavior is not yet clear, and complementary experiments would be
needed to fully understand this observation.
89
Figure 6-7: Mass-mobility plot of a 1:1 mixture of BSA digest:Spp. The band
corresponding to Spp is shown. An ion displays a shift in the drift time, as indicated by
the top white box. It matches a BSA peptide containing an oxidized methionine residue.
6.4.4. The effect of positive versus negative ionization mode
In the previous experiments, the data obtained showed that the sulfate group was
readily lost in the mass spectrometer: the DSpp peak was observed in all mass spectra
although the sample wonly contained Spp. It was reported in the literature that sulfation is
more stable in negative ionization mode for MS analyses.134 Figure 6-7 shows the
comparison of positive and negative ionization mode for the same Spp sample. In positive
ionization mode, the base peak corresponds to DSpp, which shows that the modification
is not stable under these conditions. However, in negative mode, only Spp is observed at
90
two different charge states. It was concluded that sulfation is indeed more stable in
negative ionization mode.
Figure 6-8: Analysis in positive and negative ionization mode of a Spp solution.
Negative ionization mode was a key improvement in the development of the
analytical method. It enabled collection of MS/MS data for Spp, while it was not possible
in positive ionization mode, since the slightest increase in the collision energy resulted in
the total loss of the Spp peak. Moreover, the sensitivity for Spp was dramatically
increased, as shown in Fig. 6-9: in the BSA digest. Both -1 and -2 charge states for Spp
peaks were detected, which was not the case in positive ionization. The identity of the
91
peptide was confirmed by MS/MS, leading to fragment peaks showing the modification.
The DSpp peptide was not observed.
Figure 6-9: Negative ESI-TOF MS spectrum of a 5 M BSA digest spiked with a 1:1
molar ratio of Spp.
92
Figure 6-10: MS/MS spectrum of the 1140.81 m/z (-1) peak. Enough fragments are
detected to confirm modification of a tyrosine residue by a sulfate group.
The sample was then analyzed by IMMS in negative mode (Fig. 6-11). Compared
to the same experiment in positive mode, the separation between the charge states was
more marked, and the Spp band was one of the most intense. The same behavior was
observed in negative mode regarding the drift time of the sulfated peptide: the band was
shifted to the left.
93
Figure 6-11: Negative IMMS experiment on a 5 M BSA digest spiked with a 1:1 molar
ratio of Spp.
6.5. State of the Project and Conclusion
Although not complete, the data for the analysis of tyrosine O-sulfation by IMMS
are promising. Repetition of published experiments using IMMS for screening of
phosphorylation was successful, as the expected shift in the drift time of the
phosphorylated species was observed. Notably, the shift for a peptide containing two
phosphate groups is larger than for a peptide with a single modification, due to increased
intra-molecular electrostatic interactions making the peptide structure more compact. The
sensitivity was high enough to fragment the peptides of interest with CID and collect
MS/MS data for confirmation of the sequence. The sulfated peptide could not be
identified in a BSA digest using TOF only in positive ionization mode. However, the
94
analysis of the IMMS data revealed the presence of a band shifted to the left of the
correlation line, corresponding to the sulfated peptide. The desulfated analog was not
shifted from the correlation line, showing the effect of sulfation on the drift time of
peptides. It was hypothesized that as for phosphorylation, the negative charge carried by
the sulfate group would interact with the positively charged N-terminus of the peptide
making its structure more compact. Screening of the left-shifted bands on the ion
mobilogram in negative ionization mode can lead to the identification of tyrosine Osulfated candidates, that can later be confirmed by tandem MS.
Further experiments are needed to complete this project. For tyrosine O-sulfation,
the limit of detection can be tested by using decreasing ratios of Spp in the BSA digest.
Optimized Ziptip procedure for desalting (for instance use an ion exchange column
instead of a reverse phase) could pre-concentrate the sample in highly negatively charged
peptides, giving better sensitivity for sulfated species.
95
Chapter 7: Conclusions and Future Directions
7.1. Conclusions
The work presented in this Thesis demonstrates the potential of ion mobility mass
spectrometry for the quick and easy screening for specific analysis of post-translational
modifications: SUMOylation and tyrosine O-sulfation. Although current results were
obtained from systems of low complexity, the developed methods show promising
features, such as rapid analysis, good sensitivity and selectivity for the modified peptides.
Because PTMs trigger a change in shape and charge of the target proteins, the peptides
carrying the modification have different properties than the non-modified species. IMMS
is ideal to take advantage of these features: the modified peptides can display a change in
drift time, shifting the corresponding band from the correlation lines observed in a massmobility plot. For larger modifications, such as SUMOylation, the branched isopeptides
can even carry an extra charge compared to unmodified peptides, facilitating screening
for them.
Experimentally, evidence for the SUMO isopeptides carrying a higher charge was
demonstrated with the poly-SUMO-2 and -3 systems: all of the linear peptides (but one)
were found in the 1+ and 2+ charge states, while all the isopeptides carried a 3+ charge.
Extraction of the MS spectra for the z ≥ +3 allowed for the unambiguous identification of
96
four predicted isopeptides (two produced by either trypsin or chymotrypsin cleavage, and
each – 17 Da analog). With simple model systems, the sensitivity was sufficient to obtain
data with enough resolution to accurately determine the molecular mass of the modified
species, even if the intensity of their signals were less than 35% of the base peak. The
combination of the MS and MS/MS also allowed for precise mapping of the modified
lysine peptides. Mass fingerprinting of the purchased substrates for in vitro SUMOylation
revealed that the sequence of the actual molecules did not match the supposed fragments.
Applying the method to more complex samples was hindered by this issue.
Phosphorylated casein was successfully used as a model system for generating
shifts in the drift time of modified peptides. It was also demonstrated that multiple
modifications on a single peptide increases the importance of the shift from the
correlation line. Preliminary and promising results were obtained from the extension of
the method to tyrosine O-sulfation. IMMS analysis of a solution of Spp revealed that
although the unmodified analog was found to be on the charge state correlation line as
expected, Spp displayed a shift to smaller drift times. This confirms that sulfated peptides
exhibit similar behavior in IMMS experiments as phosphopeptides. Tyrosine O-sulfation
is also likely to trigger intra-molecular electrostatic interactions due to its partial negative
charge and make the overall structure of the peptide more compact. A tyrosine O-sulfated
peptide, CCK-8 (sequence DpYMGWMDF-NH2, Spp) spiked at a 1:1 molar ratio into a
bovine serum albumin digest was detected on the mass-mobility plot, and the band
corresponding to this ion was shifted from the trend line to smaller drift time.
Experiments in negative ionization mode allowed for collection of MS/MS data for Spp,
due to the increased stability of the modification in this mode. Moreover, no product
97
resulting from the loss of the sulfate group was observed. The sensitivity is increased in
this mode, as the Spp peak could be observed in an MS scan, which was not the case in
positive ionization. Negative ionization IMMS data showed a good separation of the
different charge states, and the characteristic shift to smaller drift time for Spp was still
observed.
7.2. Future Directions
Further experiments should be conducted to complete the optimization of the
methods described in this thesis. The influence of the ionization mode (positive or
negative) should be further investigated. Negative ionization is known for generating
better signal to noise ratios for negatively charged species, which could lower the limit of
detection for the method. Once these steps are achieved the complexity of the sample will
be increased to demonstrate the utility of IMMS and determine how low the limit of
detection can be achieved.
The SUMO project, which is already at a more advanced state, can also be
improved by additional experiments. First, the method can be tested against a pool of
digested proteins containing only one SUMO substrate. Next, whole cell lysate from
human cells could be used to determine the robustness of the procedure. The current
results appear to indicate that an increased sample complexity could lead to an inability to
detect the SUMO isopeptides, especially if their abundance is extremely low. In this case,
a LC separation step might be necessary prior to MS analysis of the peptides. Another
approach relies on the fact that the addition of a second N-terminus on the isopeptides due
to the presence of the tag can also be taken advantage of: chemical modification targeting
98
N-terminal regions of peptides could be carried out and provide a more significant
difference in the gas phase behavior of modified peptides compared to the linear ones.
Furthermore, the specificity of the neutral 17 Da loss due to the N-terminal glutamine
residue that takes place for all isopeptides should be investigated as an additional
screening method for SUMO-modified isopeptides.
99
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titanium dioxide microcolumns. Mol. Cell Proteom. 2005, 4, 873-886.
134. Drake, S. K.; Hortin, G. L., Improved detection of intact tyrosine sulfatecontaining peptides by matrix-assisted laser desorption/ionization time-of-flight mass
spectrometry in linear negative ion mode. Int. J. Biochem. Cell Biol. 2010, 42, 174-179.
110
Appendix A
Tandem MS of the linear SUMO-2 peptides
Figure A-1: MS/MS of the linear SUMO-2 peptide 535.26 (+2). The confirmed peptide
is TENNDHINL. The peaks labeled with a red star (*) correspond to the neutral loss of
NH3 (− 17 Da) from a b fragment.
111
Figure A-2: MS/MS of the linear SUMO-2 peptide 599.35 (+2). The confirmed peptide
is TENNDHINLK.
Figure A-3: MS/MS of the linear SUMO-2 peptide 617.84 (+2). The confirmed peptide
is VAGQDGSVVQFK. The peaks with blue labels correspond to fragments from the
isobaric peptide KVAGQDGSVVQF.
112
Figure A-4: MS/MS of the linear SUMO-2 peptide 749.87 (+2). The only possible
peptide for the partial sequence obtained and precursor mass is DGQPINETDTPAQL.
Figure A-5: MS/MS of the linear SUMO-2 peptide 823.41 (+2). The only possible
peptide for the partial sequence obtained and precursor mass is FDGQPINETDTPAQL.
113
Figure A-6: MS/MS of the linear SUMO-2 peptide 901.46 (+2). The only possible
peptide for the partial sequence obtained and precursor mass is RFDGQPINETDTPAQL.
Figure A-7: MS/MS of the linear SUMO-2 peptide 1106.58 (+1). The only possible
peptide for the partial sequence obtained and precursor mass is VAGQDGSVVQF. The
peaks labeled with a red star (*) or a green circle (°) correspond to the neutral loss of
respectively NH3 (− 17 Da) or H2O (− 18 Da) from a b fragment.
114
Figure A-7: MS/MS of the linear SUMO-2 peptide 1342.57 (+1). The only possible
peptide for the partial sequence obtained and precursor mass is EMEDEDTIDVF.
115
Appendix B
Tandem MS simulated distribution of SUMO-2
isopeptides
Figure B-1: MS/MS spectrum of the – 17 Da trypsin SUMO-2 isopeptide (peptide 2 in
Table 5.2). The b’n fragments originating from the SUMO tag were clearly identified.
116
Figure B-2: Comparison of the experimental (blue solid trace) and simulated (red dashed
trace) isotopic distributions for isopeptide 1 in Table 5.2
Figure B-3: Comparison of the experimental (blue solid trace) and simulated (red dashed
trace) isotopic distributions for peptide 3 in Table 5.2
117
Figure B-4: Comparison of the actual (blue solid trace) and simulated (red dashed trace)
isotopic distributions for peptide 4 in Table 5.2
118
Appendix C
ESI-MS and MALDI-MS data for Sp100 and RanGAP1
fragments
Figure C-1: MS scan of the in vitro SUMOylation of the GST-tagged Sp100 fragment by
SUMO-1 in denaturing solution. No peak was detected for the Sp100 peptides or
modified peptides.
119
Figure C-2: MALDI-MS of the trypsin digested GST-tagged Sp100 commercial
fragment. MASCOT search gave a significant hit for the GST tag. Manual assignment of
the experimental peaks did not lead to matching of any Sp100 peptides.
120
Figure C-3: MS scan of the in vitro SUMOylation of the GST-tagged RanGAP1
fragment by SUMO-1 in denaturing solution. No peak was detected for the RanGAP1
peptides, modified peptides or GST tag peptides.
121
Figure C-4: MALDI-MS of the trypsin digested GST-tagged RanGAP1 commercial
fragment. MASCOT search did not give any significant hit. Manual assignment of the
experimental peaks did not lead to matching of any RanGAP1 peptides.
122
Appendix D
-casein peptides identified
Table D.1: List of the major peptides detected matching the masses of an in silico
generated list of -casein peptides. These peptides are indicated on the mass spectrum in
Figure 6-1. m/z: experimentally measured m/z value of peptide ion; z: charge of peptide
ion; Mcalculated: neutral mass of peptide calculated from the monoisotopic experimentally
measured m/z value; Mtheoretical: neutral mass of peptide calculated from sequence; ΔM:
monoisotopic mass error of measurement. MS/MS data for the confirmed peptides are
presented in Figures D-1 to D-5.
#
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
Sequence
Position
80-83
135-139
52-57
209-214
99-105
140-147
106-115
95-105
38-49
121-134
121-134
23-37
58-73
119-134
148-166
m/z
Mcalculated
Mtheoretical
M
525.3178
615.3276
689.3938
748.3659
831.3517
910.4473
1267.6862
1337.6675
1384.713
1580.8058
1660.7444
1759.9032
1927.6322
1951.8817
2316.0415
525.3178
615.3276
689.3938
748.3659
831.3517
910.4473
1267.6862
1337.6675
1384.713
1580.8058
1660.7444
1759.9032
1927.6322
1951.8817
2316.0415
525.3144
615.3283
689.3828
748.3698
831.3843
910.4741
1267.7045
1337.6808
1384.73
1580.8279
1660.7942
1759.945
1927.6916
1951.9525
2316.1369
0.0034
0.0007
0.011
0.0039
0.0326
0.0268
0.0183
0.0133
0.017
0.0221
0.0498
0.0418
0.0594
0.0708
0.0954
123
Modification
site(s)
S130
S61. S63
S130
Figure D-1: MS/MS of the -casein peptide 615.32 (+1). The confirmed peptide is
LHSMK
Figure D-2: MS/MS of the -casein peptide 748.37 (+1). The confirmed peptide is
TTMPLW
124
Figure D-3: MS/MS of the -casein peptide 1267.69 (+1). The confirmed peptide is
YLGYLEQLLR
Figure D-4: MS/MS of the -casein peptide 1384.73 (+1). The confirmed peptide is
FFVAPFPEVFGK
125
Figure D-5: MS/MS of the -casein peptide 1759.90 (+1). The confirmed peptide is
HQGLPQEVLNENLLR
126
Appendix E
BSA peptides identified
Table B.1: List of the major peptides detected matching the masses of an in silico
generated list of bovine serumalbumin peptides. These peptides are indicated on the mass
spectrum in Figure 6-4. MS/MS data for the confirmed peptides are presented in Figures
B-1 to B-15.
#
1
2
3
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
Sequence
position
281-285
101-105
205-209
205-209
490-495
236-241
29-34
257-263
249-256
161-167
37-44
549-557
548-557
66-75
35-44
402-412
421-433
438-451
347-359
437-451
m/z
z
Mcalculated
Mtheoretical
M
517.1953
545.235
649.2125
649.2125
660.2311
689.2383
712.2415
789.3269
922.3131
927.325
974.2853
1014.4284
1142.4969
1163.416
1249.3939
1305.4741
1479.5266
1511.5498
1567.4448
1639.6365
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
1
517.1953
545.235
649.2125
649.2125
660.2311
689.2383
712.2415
789.3269
922.3131
927.325
974.2853
1014.4284
1142.4969
1163.416
1249.3939
1305.4741
1479.5266
1511.5498
1567.4448
1639.6365
517.298
545.3406
649.3338
649.3338
660.3563
689.3729
712.3737
789.4716
922.488
927.4934
974.4578
1014.6194
1142.7143
1163.6307
1249.6121
1305.7161
1479.7954
1511.8428
1567.7427
1639.9377
0.1027
0.1056
0.1213
0.1213
0.1252
0.1346
0.1322
0.1447
0.1749
0.1684
0.1725
0.191
0.2174
0.2147
0.2182
0.242
0.2688
0.293
0.2979
0.3012
127
Figure E-1: MS/MS of the BSA peptide 517.34 (+1). The confirmed peptide is ADLAK
Figure E-2: MS/MS of the BSA peptide 545.38 (+1). The confirmed peptide is VASLR
128
Figure E-3: MS/MS of the BSA peptide 649.38 (+1). The confirmed peptide is IETMR
Figure E-4: MS/MS of the BSA peptide 660.40 (+1). The confirmed peptide is TPVSEK
129
Figure E-5: MS/MS of the BSA peptide 689.43 (+1). The confirmed peptide is
AWSVAR
Figure E-6: MS/MS of the BSA peptide 712.42 (+1). The confirmed peptide is SEIAHR
130
Figure E-7: MS/MS of the BSA peptide 789.53 (+1). The confirmed peptide is
LVTDLTK
Figure E-8: MS/MS of the BSA peptide 922.56 (+1). The confirmed peptide is
AEFVEVTK
131
Figure E-9: MS/MS of the BSA peptide 927.57 (+1). The confirmed sequence is
…YEIAR. The only possible peptide for the precursor mass is YLYEIAR
Figure E-10: MS/MS of the BSA peptide 974.53 (+1). The confirmed peptide is
DLGEEHFK
132
Figure E-11: MS/MS of the BSA peptide 1014.71 (+1). The confirmed peptide is
QTALVELLK. The peaks labeled with a red star (*) correspond to the neutral loss of
NH3 (− 17Da) from a b fragment.
Figure E-12: MS/MS of the BSA peptide 1163.73 (+1). The confirmed peptide is
LVNELTEFAK
133
Figure E-13: MS/MS of the BSA peptide 1305.82 (+1). The confirmed peptide is
HLVDEPQNLIK
Figure E-14: MS/MS of the BSA peptide 1479.93 (+1). The precursor ion mass and the y
fragments obtained point to the peptide LGEYGFQNALIVR
134
Figure E-15: MS/MS of the BSA peptide 1567.89 (+1). The confirmed peptide is
DAFLGSFLYEYSR
135