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Introduction to proteomics 
Basic introduction to biological mass spectrometry and proteomics 
 
 
By 
Martin R. Larsen 
 
Nicolai Bache 
Tine E. Thingholm 
Anne Kjærgaard Callesen 
Kasper Engholm-Keller 
Morten Thaysen-Andersen 
 
 
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Overview of Proteomics textbook 
 
1. Introduction to proteomics 
 
2. Biological mass spectrometry 
a. Introduction 
b. MALDI – MS 
c. ESI – MS 
d. Tandem MSMS – fragmentation and sequencing of peptides 
e. Hybride instruments 
 
3. Identification of proteins in proteomics 
 
4. Separation methods in proteomics 
a. Protein level 
b. Peptide level 
c. Affinity purification 
 
5. Quantitation in mass spectrometric based proteomics 
a. Label-free quantitation 
b. Absolute quantitation 
c. Chemical isotope labelling 
d. Metabolic labeling 
 
6. Post-translational modifications (PTMs) 
a. Phosphorylation 
b. Glycosylation 
 
7. Clinical proteomics 
a. Introduction 
b. Strategies 
c. Applications 
 
 
 
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Introduction to proteomics 
1. Proteomics 
In the past decades one of the major focuses of the biological science has been gene technology, 
especially in the field of DNA sequencing and micro-array analysis. Such interests have resulted in an 
exponential increase in the development of techniques for automated high throughput nucleic acid 
sequencing. Within the same time frame, research projects have developed in complexity from the 
characterization of one gene at a time to systematically identifying the entire genetic component of an 
organism. Complete genome sequencing of some unicellular organisms were completed already in the 
late 1970´s (e.g., Sanger F et al., J. Mol. Biol., (1978) 125(2), 225-46). The genome of the first 
eukaryotic organism (Saccharomyces Cerevisiae) was sequenced in 1996 (Goffeau A et al., Science 
1996, 274(5287), 546, 563-7), closely followed by the sequencing of the first multicellular eukaryote 
Caenorhaditis elegans in 1998 (The C. elegans Sequencing Consortium, Science 1998, 282(5396), 
2012-18) and the fruit fly, Drosophila melanogaster (Adams MD et al., Science, 2000, 287(5461), 
2185-96). Today it is routine to sequence entire genomes from your favorite organism and more than 
80 Eukaryotic genomes has been sequenced, including the Human genome which was completed in 
2006 (Gregory SG, Barlow KF, McLay KE, et al., Nature, 2006, 441 (7091): 315–21). The haploid 
human genome contains an estimated 23,000 protein-coding genes, far fewer than had been expected 
before its sequencing (International Human Genome Sequencing Consortium. Nature, 2004, 431 
(7011): 931–45). In fact, only about 1.5% of the genome codes for proteins, while the rest consists of 
RNA genes, regulatory sequences, intron and (controversially) "junk" DNA (International Human 
Genome Sequencing Consortium Nature, 2001, 409 (6822): 860–921). 
The genome defines a delicate framework of highly complex intracellular processes within a cell. Even 
though genetic information provides the basic scaffold for life, there is no direct correlation between 
genomic information and the proteins present in the cell and thereby the phenotype of a living cell or 
organism. A beautiful example of this higher order of complexity is the amazing transformation of a 
caterpillar into a butterfly (see Figure 1.1). They are genetically identical but have two completely 
different phenotypes and from the figure a significant change in the expressed proteins is obvious 
represented by two dimensional gel electrophoresis. In between these phenotypes a large number of 
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transition states exist, each with a different subset of proteins expressed at the given transition state. 
Another interesting fact is that the identity between the genome of chimpanzee and humans is 98.5% 
indicating a significant similarity; however, the phenotypes are completely different. The differences 
observed at the phenotypic level cannot be accounted for by the genomic differences alone. Thus 
differences between cellular states and cellular function are reflected in differential gene expression 
being manifested at both the mRNA and protein level. However, there are no direct correlation between 
intercellular mRNA levels and protein concentration due to intensive regulatory steps at both the 
mRNA and protein level (Anderson et al. 1997; Steiner et al. 2000; Hegde et al. 2003). Thus protein 
abundance in a biological system is essentially dependent on the rate of protein transcription vs. 
degradation and can only be measured directly at the protein level and not on the level of the mRNA. 
This is true for the majority of the proteins in the cell however, housekeeping proteins such as actin, 
keratin, tubulin have a very high correlation between mRNA and protein. 
 
 
 
 
 
 
 
 
 
 
Figure 1.1: The transformation of a caterpillar into a butterfly. The 2D gels show the different 
protein pattern present in the different steps during the morphological changes. 
 
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Additional complexity and diversity of the cell is introduced by post-translational modifications (PTM) 
of proteins (i.e., modifications of proteins introduced after the protein has been synthesized). These 
modifications are often chemical groups which are covalently linked to the proteins by the action of 
specific enzymes, e.g. disulfide linkage, glycosylation and phosphorylation. Some of the PTMs are 
reversible modifications e.g., phosphorylation, whereas some are irreversible, e.g., glycosylation. In 
total more than 200 PTMs are known today and many more are likely to exist in nature. PTMs have 
been shown to be extremely important in stabilising and influencing protein structure, defining 
localisation of proteins e.g., via lipid anchors, or they can act as key regulator in diverse cellular 
processes such as metabolic pathways, signal transduction, cell growth, etc. (Hunter 1998; Cohen 2000; 
Hunter 2000; Cohen 2002). Thus, regulation of protein expression and PTMs is directly responsible for 
virtually all cellular functions. 
The human genome has revealed about 23000 coding sequences (proteins) however due to post-
transcriptional processing (splicing) and post-translational processing (PTMs, specific proteolysis etc.) 
the true amount of protein species in a living cell could be in the area of several hundred thousand (see 
Figure 1.2), which puts high demands for any analytical technique to analyze the whole subset of 
proteins – the proteome - of a living cell. 
The term “proteome” was invented in Sydney, Australia, in 1995 and is defined as the entire 
complement of proteins expressed by any given genome, cell, tissue or organism (Wasinger et al. 1995; 
Wilkins et al. 1996) at a given time, and the term proteomics is the large scale study of proteomes. 
More refined definitions have later been applied and today if one is working with a subset of proteins 
they often call this proteomics too, e.g., the mitochondrial proteome, plasma membrane proteome etc. 
The key to proteomics is the possibility to identify and quantify proteins in complex samples in 
different situations, e.g., healthy versus disease or control versus drug treated cells. As such, 
protoemics is very technological minded. In order to achieve identification and quantitation of proteins, 
a large number of strategies have been developed over the last decade. In the beginning of the 
proteomic field (prior to 2000) most of the proteomics work was performed using two-dimensional gel 
electrophoresis (2DE) to separate proteins from each other (see more details in Chapter 4.1) and then 
combine this technique with a sensitive way to identify the proteins in thegels. The latter was initially 
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performed using technologies like traditional Edman amino acid sequencing and amino acid analysis 
combined with sophisticated database search software programs. However, these methods are not very 
sensitive, with detection limits in the low pico-mole area at best, and they are therefore not able to 
identify proteins present in small amount on the gel. Thus, only very abundant proteins could be 
identified. 
With the introduction of very robust biological mass spectrometry (MS) in the late 1990 it became 
possible to use MS to identify and quantify proteins from both gels and solution samples. Today most 
protein identification and quantitation in proteomics is performed using MS technologies. 
 
 
 
 
 
 
 
 
 
 
 
Figure 1.2: Illustration of the process from gene to gene product. The gene is transcribed into a 
primary RNA product by protein complexes and thereafter the introns are spliced out using 
specific enzymes giving mRNA molecules. The latter process can in many eukaryotic cells take 
place in different ways giving raise to many mRNA molecules from a single gene. This process 
is called alternative splicing. After translation of the mRNA the proteins can become modified 
 
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by various chemical groups (post translational modifications). The number of gene products can 
be several thousands in a single cell. 
Traditionally proteomics can be divided into three main areas: expression proteomics, modification 
specific proteomics (modificomics) and interactomics. 
Expression proteomics involves the identification and quantitative analysis of proteins at their 
expression level. This means identification of which proteins are present in a given proteome at a given 
time and in want amount relative to the other proteins. The classical approach to address these 
questions is by a comparative gel-based proteomics strategy, see Figure 1.3. Two or more proteomes 
can be compared by two-dimensional gel electrophoresis (2DE) which separate proteins according to 
isoelectric point and molecular weight, and the individual spots, representing individual protein 
isoforms, can be relatively quantified by spot density, using specialised imaging software. The 
individual spots are excised from the gel and the protein is in-gel digested by a proteolytic enzyme. 
Peptides are subsequently extracted from the gel piece and they are analyzed by MS to obtain very 
accurate masses of each peptide. The mass of the peptides can then be used as a fingerprint to identify 
the protein in protein sequence databases (Shevchenko et al. 1996). The 2DE gel-based approach is 
very laborious and have several inherent shortcomings, such as decreased analysis of low and high 
molecular weight proteins, hydrophobic proteins and various modified proteins (Gygi et al. 2000). 
However, specialized buffers and various techniques can overcome some of these problems, including 
the application of very strong detergents in the IEF loading buffer and specialized gels. 
The second strategy is MS-based and is frequently referred to as liquid chromatography (LC)–MS. This 
technique is becoming increasingly popular due to the significantly lower amount of material used for 
the analysis and the direct sequencing of peptides for identification. Here the proteins in a solution are 
processed into peptides using specific proteases and then the peptides are sequenced and identified by 
MS techniques and bioinformatics. However, the main limitation of LC-MS is the huge amount of 
peptides that can be present in a complex protein mixture such as a whole cell lysate. For example if we 
look at the yeast proteome it is expected to include about 6300 proteins which after proteolysis will 
result in > 350.000 peptides. Despite days of LC-MS analysis of the yeast proteome only a subset of 
the expressed proteins has until today been identified with this approach. If we then look at more 
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complex eukaryotes like human the picture get even more complex. If we only had 23000 gene 
products in human cells we will after processing using a specific protease end up with > 2 million 
peptides which had to be sequenced by LC-MSMS. This is a significant challenge to any analytical 
technique and therefore not readily achievable with the present technology. 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 3: Strategies for expression proteomics. Expression proteomics can be divided into a 
gel-based strategy and a MS driven strategy. See text for further details. 
For the quantification of proteins in LC-MS approaches, a chemical reagent can be introduced onto an 
amino acid functional group (e.g., primary amine), either prior to enzymatic processing or after (see 
later under quantification). Briefly, if two samples are to be compared, the first sample is labelled with 
a light isotopic version of the chemical reagent, and the second sample is labelled with a heavy isotope 
of the same chemical regent. The labels can be introduced on the protein or peptide level depending on 
 
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the individual strategy. After the labelling and processing of the samples, they are consolidated, and the 
peptides are analysed by multi-dimensional LC/LC-MS and MS/MS to facilitate protein identification 
and quantification. New technology and advances in instrumentation with increased resolution have 
improved this strategy, and it is now becoming the dominating workflow in proteomics. 
 “Modificomics” deals with large-scale identification, characterization and quantification of PTMs on 
proteins in the cell. Virtually all cellular processes are tightly regulated by proteins with distinct 
functions. One of the fastest ways to alter the function of a protein is by PTMs. These can modulate 
many properties: conformational state, stability, activity, interaction partners and subcellular 
distribution. Many of the control mechanisms involved in the dynamic regulation of cellular expression 
patterns in health and disease are dependent on specific PTMs within key proteins. Therefore, the 
determination and quantitative assessment of PTMs in proteins is fundamentally important for 
elucidation of the complex processes that govern cellular events such as cell growth, division, 
differentiation and migration. Such events are implicated in disease development and progression. 
Several new and exciting protein chemistry tools have been developed to be able to identify and 
characterise PTMs, which are typically present on low abundant proteins in the cell, especially if there 
is a low stoichiometry (low occupancy of the modified site). Therefore it can be very difficult to 
characterise modified peptides together with unmodified un-modified peptides. The development of 
methods to enrich and analyse different PTM sub-proteomes i.e. the phosphoproteome, by selective 
affinity methods like immobilised metal affinity chromatography (IMAC) (Neville et al. 1997) and 
titanium dioxide (TiO2) chromatography (Pinkse et al. 2004; Larsen et al. 2005) has greatly improved 
the success and analytical depth of these workflows. Modificomics has significantly benefited from 
instrumental improvements towards sensitivity and analytical speed combined with specialised peptide 
fragmentation capabilities, e.g., MSn, multistage activation (MSA) (Schroeder et al. 2004) and electron 
transfer dissociation (ETD) (Schroeder, M. J., et al., 2005 Journal of Proteome Research 4, 1832-41). 
This improves the success of sequencing modified peptides, e.g. phosphopeptides, which otherwise 
show an all-dominating neutral loss of phosphoric acid, which makes traditional sequencing analysis 
difficult. 
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“Interactomics”involves elucidation of how proteins interact to form large functional complexes and 
characterization of their dynamical and structural properties. Protein activity is greatly affected by 
protein folding and protein-protein interactions which cannot be directly addressed in the classical 
expression proteomics. Thus it is essential to obtain a fundamental understanding of the function of 
proteins and protein complexes at the molecular level. For this purpose nuclear magnetic resonance 
(NMR), x-ray crystallography and hydrogen-deuterium exchange (1H/2H) combined with MS (HX/MS) 
are three complementary and very powerful techniques to investigate the native dynamic and structural 
properties of proteins in solution. In addition, in order to characterize protein complexes several affinity 
purification strategies has been developed to specifically purify whole complexes and study the 
different subunits using MS approaches. 
 
 
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2. Biological Mass Spectrometry 
Almost a century ago Thomson developed the very first mass spectrometer (Thomson, 1913) and in 
1918 Arthur Jeffrey Dempster made the first modern mass spectrometer. They were able to accurately 
measure the natural occurring elements and their isotopes. The first attempts to analyse peptides and 
proteins using mass spectrometry date back to the mid-1960’s where electron ionisation (EI) and gas 
chromatograph (GC) mass spectrometry were used (Barber et al. 1965; Biemann et al. 1966). The 
major drawback was that only volatile samples could be ionised and subsequently analysed by a mass 
spectrometry, limiting the technology to small apolar molecules. By contrast, peptides and proteins are 
rather large polar molecules that are not easily volatized and undergo fragmentation upon conventional 
thermal heating. In order to get large polar molecules volatile they had to be derivatized. Even though 
chemical derivatization (CI) techniques allowed a limited number of small biomolecules to be analysed 
by increasing analyte volatility, sensitivity was poor (Kiryushkin et al. 1971). At this time mass 
spectrometry was considered of little use for biological applications. But the situation was improved 
with the introduction of field desorption (FD) allowing un-derivatized peptides and small proteins to be 
analysed (Winkler et al. 1972). However, sensitivity was low, and the technique was demanding and 
required an experienced operator. A milestone was reached with the introduction of plasma desorption 
(PD) (Torgerson et al. 1974) and fast atom bombardment (FAB) (Barber et al. 1981) mass spectrometry 
in the early 1980’s making mass spectrometry a viable tool for the analysis of larger and thermally 
labile biomolecules such as peptides and proteins. The decisive breakthrough in the analysis of larger 
proteins and peptides came with the development of MALDI by Michael Karas and Franz Hillenkamp 
(Karas et al. 1987; Karas et al. 1988) and Electrospray ionisation by John B. Fenn (Fenn et al. 1989). 
These soft ionisation techniques have revolutionized protein mass spectrometry by allowing high-
molecular-weight compounds to be analysed as intact molecular ions with very little fragmentation. 
The scientific impact of these achievements was acknowledged in 2002 and awarded the Nobel Prize in 
Chemistry. The prize was shared between scientists working with MS and nuclear magnetic resonance 
(NMR). Koichi Tanaka and John B. Fenn shared the price in MS for the development of soft laser 
desorption (SLD) and ESI, respectively. However, this is very controversial and many scientists in the 
MS world believe that Michael Karas and Franz Hillenkamp should have been awarded the Nobel prise 
for the development of MALDI instead of Koichi Tanaka. They described the concept of matrix-
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assisted laser desorption (MALD) (Karas et al. 1985) and later the now widely used solid-phase 
MALDI technique used in modern MALDI mass spectrometers (Karas et al. 1988). The soft laser 
desorption (SLD) method developed by Koichi Tanaka was based on a liquid matrix consisting of a 
mixture of analyte and glycerol with ultra fine cobalt particles as a matrix (Tanaka et al. 1988). This 
technique has not been used in the field of protein mass spectrometry. MALDI quickly became the 
dominating ionisation technique in the field of proteomics together with ESI. 
2.1. Matrix-assisted Laser Desorption/Ionisation 
The early experiments with laser desorption date back to 1960 (Vastola et al. 1970), and the first 
systematic attempts to generate ions of organic molecules with lasers date to the early 1970s 
(Posthumus et al. 1978; Kupka et al. 1980). However, all experiments on laser desorption of organic 
ions revealed an upper limit of approximately 1000 Da for biopolymers that could be desorped as intact 
ions. However, this limit was dependent on the molecular structure and laser parameters, also making 
reproducibility very difficult. The situation changed dramatically with the development of MALDI by 
Hillenkamp and Karas in 1985-1988 (Karas et al. 1985; Karas et al. 1987; Karas et al. 1988). This 
method increased sensitivity, and because the process is independent of the absorption properties and 
size of the compound being analysed, it allows very big molecules to be desorped, ionised and 
subsequently analysed. 
 
Figure 2.1: Ion formation by MALDI. 
Illustration of the principle of MALDI 
from irradiation to ion formation by the 
gas-phase proton transfer model. For 
further description of the MALDI process 
see the text. 
 
 
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The first step in MALDI is to create a “solid solution” deposit of analyte-doped matrix crystals, where 
the analyte molecules are embedded throughout a matrix so they are completely isolated from one 
another. Matrix is normally a small acidic molecule. In doing so, the matrix is believed to serve two 
major functions: 1) absorption of energy from the laser pulse, which minimizes sample damage by 
absorbing most of the incident energy and increases the efficiency of energy transfer from the laser to 
the analyte, and 2) isolation of the analyte molecules from each other preventing their interaction – 
molecular aggregation (Hillenkamp et al. 1991). When the analyte-doped matrix crystals are irradiated 
by a short laser pulse, it induces a rapid heating of the crystals by the accumulation of a large amount 
of energy through excitation of the matrix molecules. The rapid heating causes a local sublimation of 
the matrix crystals (like a local explotion in vacuum) and an expansion of the matrix into the gas phase, 
bringing intact analyte molecules into an expanding matrix plume (Dreisewerd 2003). An illustration of 
the MALDI process is shown in Figure 2.1. 
Only little internal energy is transferred to the analyte molecules and they may be cooled during the 
expansion process. Ionisation of the analyte molecules can occur at any time during this process, and 
there are several theories trying to explain the process, reviewed in (Zenobi et al. 1998; Dreisewerd 
2003; Karas et al. 2003; Knochenmuss et al. 2003). One of the most widely accepted theories on the 
ion formation mechanism involves a gas-phase proton transfer in the expanding plume with photo-
ionised matrix molecules (Zenobi et al. 1998), see Figure 2.1. 
Several different matrices are used in MALDI MS and there are numerous different sample preparation 
techniques. Some of the most commonly used matrices are listed in Table 1 and their chemical 
structures are shown in Figure 2.2. 
Matrix Chemical name Application 
-cyano (HCCA) -Cyano-4-hydroxycinnamic 
acid 
Peptides, proteins and organic 
compounds 
Sinapic acid (SA) 3,5-Dimethoxy-4-
hydroxycinnamic acidHigher mass biopolymers 
Gentisic acid (DHB) 2,5-Dihydroxybenzoic acid Peptides, proteins and 
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 carbonhydrates 
PA Picolinic acid Oligonucleotides 
HPA or 3HPA 3-Hydroxypicolinic acid Oligonucleotides 
Trihydroxyacetophenone 2,4,6-Trihydroxyacetophenone Oligonucleotides, peptides 
ATT 6-Aza-2-thiothymine Oligonucleotides 
 
Table 1: The matrices most commonly used for MALDI along with their chemical names and 
applications are listed in this table. 
COOH
OH
HO
DHB
HO
COOH
CN
4-Hydroxy--cyanocinnamic 
acid (4HCCA)
HO
OCH3
CH3O COOH
Sinapinic acid (SA)
COOH
N
Picolinic acid (PA)
COOH
N
HO
3-Hydroxypicolinic acid (3HPA)
OH
OHHO
CH3
2,4,6.Trihydroxy 
acetophenone
N
N
HN
HS
O
CH3
6-Aza-2-thiothymine (ATT)
O
 
Figure 2.2: The chemical structures of the matrices listed in Table 1. 
Matrices are classified as either “hot” or “cold” matrices depending on the degree of fragmentation 
they cause to the sample upon laser desorption. DHB does not give rise to much fragmentation and is 
therefore considered to be a cold matrix, whereas -Cyano-4-hydroxycinnamic acid is a hot matrix. 
Several matrix additives have been shown to improve the efficiency of the ionization of difficult 
analyte molecules in MALDI MS. When analyzing phosphopeptides by MALDI MS the addition of the 
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mineral acid ortho-phosphoric acid (H3PO4, PA) to the DHB matrix has shown to improve the signal of 
phosphopeptides in particularly for multiphosphorylated peptides (Kjellstrom and Jensen, 2004). In 
addition, ammonium citrate has been shown to improve signals from phosphopeptides/proteins in 
MALDI MS (Asara and Allison 1999). The exact mechanisms behind the addition of additives are not 
known, however it is believed that additive can change the physionchemical properties or both the 
matrix or the analyte molecules and thereby change solubility and incorboration in crystals. 
MALDI can practically be coupled to any mass analyzer, but it is typically combined with time-of-
flight (TOF) analyzers. MALDI is an ion source that generates pulsed ion packages due to the laser 
desorption process and is for that reason very compatible with TOF mass analyzers and mass analyzers 
that allow the storage of ions such as ion trap and FT instruments (see later). 
Some of the obvious advantages using MALDI MS are the rapid sample preparation and analysis (one 
analysis can take less than a minute). It is a very sensitive method and the technique is suited for 
analysis of heterogeneous and semi-complex samples. Besides this, MALDI MS has a high tolerance 
towards salt and other low molecular contaminants making it ideal for analysis of peptides or proteins 
from biological samples. One disadvantage of MALDI is that the use of matrix gives rise to matrix 
background in the spectra below 1000 Da, which can suppress the signals from compounds in the low-
molecular-weight range. This interference varies between different matrices. DHB is good for small 
molecules and peptides (200-1000 Da), since it creates minimal amount of interference in this area. 
Another disadvantage with MALDI is that sequence information by ion fragmentation using this ion 
source relies on singly charged ions, which require much more energy (Hoffmann and Stroobant, 2001) 
and is significantly dependent on the amino acid sequence of the peptide to be analyzed. 
Recently, MALDI has been coupled with offline HPLC using robots, which deposit small fraction of 
the eluted peptides directly onto the MALDI target (Mirgorodskaya, et al., 2005). This has some 
advantages over HPLC coupled to ESI (see later), since the entire sample is not used for spraying, 
however, this is not routine in many laboratories. 
 
 
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2.2. Electrospray ionisation 
Electrospray ionisation (ESI) is a versatile and commonly used method for the production of gas phase 
ions from a liquid. The ion source operates at atmospheric pressure and produces a continuous flow of 
quasimolecular ions ideal for several state-of-the-art MS configurations. The pioneering experiments in 
1917 on Electrospray ionisation (ESI) by the physicist John Zeleny precede the first description of the 
process by Malcolm Dole in 1968 (Dole et al. 1968). The well-defined breakthrough and introduction 
of electrospray ionisation into a useful technique for mass spectrometry of large biomolecules was 
presented by John Fenn in 1988 at the 36th ASMS conference in San Francisco and published in 1989 
(Fenn et al. 1989). Later it was optimised into nanoflow ESI (nESI) with increased sensitivity by Wilm 
and Mann (Wilm et al. 1996). The nESI source is now routinely used in front of most state-of-the-art 
MS configurations and is well suited for solvent based protein and peptide separation techniques like 
high pressure liquid chromatography (HPLC) and nanoflow HPLC ideal for high throughput nano 
HPLC-MS applications. 
Electrospray ionisation works by introducing the sample solution into the source region of the mass 
spectrometer through a small steel or glass capillary. By applying a high potential (±1700-5000 v) 
between the capillary and the counter electrode (orifice), a strong electric field is created. This induces 
a charge accumulation at the liquid surface at the tip of the capillary (Kebarle 2000), see Figure 2.3. 
The ions will be drawn from the capillary and attracted towards the counter electrode. However, the 
surface tension of the liquid provides an opposing force holding the liquid together which results in the 
formation of a Taylor cone at the tip of the capillary. As the charge density builds up at the tip of the 
Tayler cone, the repulsive electrostatic forces will overcome the surface tension and produce a jet of 
charged droplets, a process known as budding. The charged microdroplets undergo evaporation as they 
progress down a pressure gradient in the ion source reducing their size. Consequently the charge 
density builds up until the Rayleigh limit is reached. At this point the accumulated repulsive coulombic 
forces exceed the surface tension of the microdroplet and a coulomb explosion (droplet fission) occurs. 
The charge to mass ratio (of droplet) is increased in the now smaller daughter droplets relative to the 
parent droplet. However, the increased charge at the surface is balanced by the increased surface-to-
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volume ratio of the daughter droplet. Upon several consecutive rounds of coulombic fission very small 
charged droplets are formed (Kebarle 2000). 
There are two major competing theories for the final ionisation step of the analyte molecules. The 
charged residue model (CRM) first suggested by Dole et al. (Dole et al. 1968) and the ion evaporation 
mechanism (IEM) based upon the ideas of Iribarne and Thompson (Iribarne et al. 1976; Thomson et al. 
1979). Both are illustrated in Figure 2.4. The CRM model suggests that charged droplets undergo 
several consecutive rounds of coulombic fission and eventually produce droplets containing a single 
analyte molecule. Upon complete solvent evaporation, the molecule is left with the charge carried by 
this droplet (Schmelzeisen-Redeker et al. 1989). The IEM model suggests that as the droplet reaches a 
certain radius the ions are desorbed directly out of the droplet by the strong electric field at the surface 
of the droplet (Thomson et al. 1979). It is most likely a mixture of the two ionisation mechanisms that 
account for the ionisation in ESI. 
 
 
 
 
 
 
 
 
Figure 2.3. Ionisation of proteins and peptides by electrospray ionisation. At the tip of the 
capillary/needle, the liquid forms a Taylor cone. At the tip, small droplets arebudding off, 
forming small charged droplets. As the liquid evaporates the droplets undergo several rounds of 
coulombic fission and eventually form individual analyte ions on the gas phase. See the text for 
further details. Adapted and modified from (Hoffmann et al. 2001) 
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Figure 2.4. Figure illustrating the two different theories for generating molecular ions in ESI. 
ESI normally produces multiply charged ions, and this allows molecules that have masses exceeding 
100 kDa to be analysed by mass spectrometers with low mass/charge (m/z) range analysers. A standard 
quadrupole analyser has an upper m/z limit of approximately 4000 (Hoffmann et al. 2001). In the 
majority of MS driven proteomic workflows, peptides of approximately 600 – 3500 Da are analysed 
producing mainly 2+, 3+ and 4+ ions with ESI. These multi-charged ions are well suited for collision-
induced dissociation (CID) which is more efficient and requires less collision energy compared to CID 
of singly charged ion (Wysocki et al. 2000). The same is true for electron capture dissociation (ECD) 
and electron transfer dissociation (ETD) where a charge state of 3+ and 4+ is desirable for optimal 
fragmentation efficiency (Zubarev et al. 1998; Mikesh et al. 2006)(see later). 
In positive ion mode the analyte molecules are sprayed at low pH to improve the formation of positive 
ions, whereas higher pH conditions are used for negative ion mode. Proteins and peptides are usually 
analyzed using positive ion mode, whereas oligonucleotides and oligosaccharides are analyzed using 
negative ion mode. The number of positive charges obtained by a molecule is related to the number of 
basic sites on the molecule, however, posttranslational modifications also influence this. 
 
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In the electrospray ionization process, neutral species, ions and clusters of ions with neutrals are 
formed. The ions need to be separated from the neutrals to aid desolvation before entering the mass 
spectrometer. This can be accomplished simply by positioning the capillary slightly off-axis from the 
entrance to the mass analyzer, since the outer regions of the spray consists mainly of smaller, lighter 
and more desolvated droplets (Cech and Enke, 2001). Another important problem is the cooling created 
by the analyte and the solvent adiabatic expansion, which produces ion clusters. This is prevented by 
the application of a heated, dry nitrogen counter-current or gas curtain. 
There are several advantages using electrospray as an ion source. As mentioned above multiply 
charged ions are produced, which makes the analysis of high-mass ions by mass analyzers with relative 
low m/z range possible, in addition to giving a better mass accuracy. In addition, fragmentation of 
multiply charged peptides requires less energy and is therefore easier. No matrix is needed so no 
signals from matrix will interfere with the analyte signals and in principle electrospray is unlimited in 
mass range (Siuzdak, 1996). However, multiply charged ions are a disadvantage for low resolution 
instruments such as ion traps as no charge stages can be deduced. In addition, the ESI system has a low 
salt tolerance and the purity of the sample needs to be fairly high. Compared to the MALDI techniques 
there is a great loss of sample. The electrospray ion source is compatible with practically any type of 
mass spectrometers. Most commonly used instruments are the quadrupole and the ion trap (Mann and 
Wilm, 1995). The coupling of ESI with LC (LC-ESI) have become fully automated as opposed to LC-
MALDI, which is only semi-automated. 
 
2.3 Mass analyzers used in modern biological mass spectrometry 
Mass spectrometers do not actually measure mass, but rather the mass-to-charge (m/z) ratio of ions, 
expressed in units of Thomson (Th) or atomic mass units (u) for singly charged ions (z = 1). As there 
are many different ion sources, there are a variety of mass analyzers, each working by different or 
similar mechanisms in order to determine the m/z ratio of ions. Some mass analyzers scan the ions 
formed in the ion source, while others have the capability to isolate individual ions due to their m/z 
ratio. 
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Four important factors come into play, when describing mass analyzers. 
1. The upper mass limit, 
2. The transmission, 
3. The resolution (R) 
4. Mass accuracy. 
The upper mass limit is the highest possible m/z ratio that the particular instrument is capable of 
measuring. The transmission describes the number of ions reaching the detector compared to those 
produced by the ion source. A low transmission indicates a substantial loss of ions from the ion source 
to the detector and results in a decrease in sensitivity. Normally there is a significant loss of ions from 
the ion source to the mass analyzer in any mass spectrometer. The resolution is a term for how well the 
instrument is able to distinguish between signals from two ions of similar m/z ratios. 
The definition of the resolution R is given by the equation: 
 R = m/m 
Where m is the smallest mass difference between two peaks with masses m and m + m, respectively. 
The resolution can also be determined from one peak with mass m as the peak width m at x% of the 
peak height, usually being 50%, designating m as the full width at half-maximum (FWHM) (Figure 
2.5). 
 
 
 
 
 
 
In
te
ns
it
y
m/z
m
x %
2x %
100 %
m
 
21 
 
Figure 2.5. Relationship between two definitions of resolution. This figure was adapted from 
Hoffman and Stroobant, 2001. 
Finally the fourth and very important factor is the mass accuracy. This is a measurement of how well a 
measured m/z ratio correlates with the real (theoretical) ratio. The mass accuracy is highly dependent 
on how well the instrument is calibrated and is therefore somewhat variable (Hoffmann and Stroobant, 
2001). The data can be recalibrated after the analysis if necessary, which will improve the mass 
accuracy. The mass accuracy is also highly influenced by the resolution. The higher the resolution, the 
better the separation of the isotopes and the more accurate peak assignment can be performed and 
consequently better mass accuracy is achieved (Figure 2.6). 
The contributions of these four factors determine the performance and sensitivity of the individual MS 
instruments. 
 
 
 
 
 
 
 
Figure 2.5: Resolution. The three sprectra were calculated for the same molecular formula at 
resolution (R) 200, 2000, and infinity () using the 10% ratio definition (10% above baseline). 
The higher the resolution is, the narrower the peaks will be. This figure was adapted from 
Siuzdak, 1996. 
 
 
R
el
at
iv
e 
in
te
ns
ity
(%
)
R = 200
R = ∞
R = 2000
m/z
 
22 
 
The Quadrupole (Q) 
The principle of the quadrupole was first described by Paul and Steinwegen in 1953 (Paul and 
Steinwegen, 1953). A quadrupole consists of four parallel rods with a circular or hyperbolic section 
(Figure 2.7). A direct current (fixed DC) and a superimposed radio-frequency (alternating RF) 
potential are applied to the rods. 
 
 
 
 
 
 
 
 
 
 
 
 
 
Figure 2.7. Skematic illustration of the principle of the quadrupole. This figure was adapted 
from Skelkton and Gates, 2000. (http://www-methods.ch.cam.ac.uk/meth/ms/theory/quadrupole.html) 
Ions enter the quadrupole in the space between the four rods and travel along the z-axis. At the same 
time ions move circularly in the x-y plan due to the alternating electric field. The opposing rods have 
the same charge, whereas the rods next to each other haveopposite charges (Figure 2.7). Positive ions 
are drawn to the negatively charged rods (- 0). Due to a change of the potential polarity, the ions are 
Nonresonant ion
Resonant ion (detected)
Quadropole rods
Source slits
(entrance to the quadropole)
To detector
Analyte ions 
(with different m/z ratios)
X
Z
Y
- 0
+ 0
r0
 
23 
 
repelled by the rods and change direction before they touch the rods and discharge themselves, which 
would prevent their detection. Instead, they are positioned in the center of the quadrupole. 
The stability of the trajectory of a given ion depends on its m/z ratio. Only ions that are stable will be 
able to pass through the quadrupole. This means that by changing the DC/RF potentials on the rods, the 
quadrupole can select ions of a particular m/z ratio. At a given time only ions of this particular m/z ratio 
are let through the quadrupole and will be scanned by the detector, whereas ions with other m/z ratio 
are unstable and will discharge themselves against the rod surfaces. This relation gives rise to the so-
called stability areas, in which ions are successfully scanned (Figure 2.8). 
 
 
 
 
 
 
 
 
Figure 2.8: The stability areas are created as a function of DC voltage (U) and RF voltage (V) 
for ions with different masses (m1 ‹ m2 ‹ m3). Making U as a linear function of V results in a 
straight line (scan line) intersecting at (0;0). Choosing U and V ratios from this line will give 
good scan of the individual masses (m). Choosing U and V ratios below the scan line will result 
in an overlap of different m ratios, decreasing the resolution. This figure was adapted from 
Hoffmann and Stroobant, 2001. 
 
As illustrated on the figure, keeping U/V constant allows for the detection of several different masses 
(along the scan line). The higher the slope of the line, while still intersecting a stability area, the better 
U
(D
C
 V
ol
ta
ge
)
V (RF Voltage)
m1
m2
m3
Unstable area
Unstable
area
(constant DC/RF)
Scan line
 
24 
 
the resolution. At U = 0, when no direct potential is applied, the resolution is zero. However, ions are 
still transmitted as long as V is within a stability area. The more the instruments are operated at the 
peaks of the figure, the better the resolution will be, but there will be some overlap between the 
different masses below the scan line and therfore the resolution will decrease. The mass range and 
ultimate resolution of a quadrupole instrument depends on the length and diameter of the rods. 
However, the mass range for common quadrupole instruments is about 4000 Th but can be extended up 
to 8000 Th and their resolution operating in normal scan mode is only approaching 3000, classifying 
them as low-resolution instruments (Hoffmann and Stroobant, 2001). An iontrap can achieve a very 
high resolution if operated in enhanced or zoom scan mode, however with a significant drop in speed. 
Quadrupole instruments can be used in tandem mode as well, where instruments are connected linearly. 
Often the quadruple instruments are coupled to a time-of-flight instruments (See section on QqTOF) or 
several quadrupoles are connected (triple quadrupole (QqQ)). 
 
Time-Of-Flight (TOF) 
Time-of-Flight mass spectrometry is one of the simplest methods of mass measurement although there 
are hidden complexities when it comes to higher resolution applications. The idea of linear time-of-
flight (TOF) mass spectrometry was introduced by Stephens in 1946 (Stephens, 1946) and some ten 
years later the design for the first TOF instrument was published by Wiley and McLaren (Wiley and 
McLaren, 1955). A TOF mass spectrometer consists of an ion source in which the ions are ionized and 
accelerated, a field-free flight tube in which the ions are separated according to their m/z ratio and a 
detector, in which the ions are detected (Figure 2.9). 
 
 
 
 
 
25 
 
 
 
 
 
 
 
 
 
Figure 2.9. Schematic presentation of the time-of-flight (TOF) principle using extraction of ion 
packages (MALDI). Ions are produced in the ion source and accelerated in a strong electric 
field before entering the field-free drift region. Finally, the ions reach the detector at different 
times depending on their velocities. Vacc being the acceleration potential for the ions and Ek the 
kinetic energy. 
After ionisation in the ion source, the ions are normally accelerated by a 20 to 25 kV potential (Vs), and 
obtain the kinetic energy of Ek = Vsq = mv
2 /2, where q is the charge of the ion, m is the mass of the ion 
and v is the velocity of the ion. Therefore the smaller ions will travel faster than the larger ions, and 
will reach the detector first. By measuring the flight time of the ions, the m/z can be calculated from t2 
= (m/z) k, where k = d2/(2Vse), d being the distance the ion travels and e the charge of an electron. 
There is no limit to the upper mass range of a TOF analyzer, which makes it highly compatible with 
soft ionization techniques where only molecular ions are produced. MALDI produced ions in pulses 
making it a good choice for coupling with TOF, since the TOF analyzer is a pulsed instrument 
requiring a well-defined starting time (point of initiation). TOF mass analyzers are also very sensitive 
instruments because of a high transmission efficiency due to the ability to detect ions over the whole 
m/z range at the same time without scanning. However, there are limitations to the simple construct of 
Detector
Positive ions
Analyzer Field-free Region
+20.kV
Ion Source
d
Laser beam
Ek = 0Ek = Vacc/dS
dS
Flight tube
sample
0.kV
 
26 
 
TOF, resulting in a poor resolution. This is due to different factors creating the distribution in flight 
times for ions of the same m/z ratio (Hoffmann and Stroobant, 2001). Some of which are: 
Kinetic energy distribution: In theory all ions are subjected to the same acceleration potential (Vacc) 
in the ion source, but in practice there will be variation in their initial kinetic energies. This will result 
in different velocities for ions of identical masses so these will not reach the detector at the same time. 
Time distribution: Variations of the length of the ion formation in the source. This leads to a minor 
time shift between different ions leaving the ion source, so these ions will not reach the detector at the 
same time. 
Space distribution: Variations of the spatial positions of identical ions in relation to the ground grid 
gives a difference in the energy acquired. The ion closest to the ground grid will not be in the energy 
field as long and therefore receive less energy. The ion with the highest kinetic energy will reach the 
detector first. 
Distribution of the ion direction: Identical ions with the same initial kinetic energy positioned at the 
same initial position, but with opposite directed directions will eventually both reach the detector, but 
not at the same time. 
Equipment: Electronics and especially the digitizers. 
The two-stage acceleration strategy (delayed extraction) developed by Wiley and McLaren in 1955 
corrects both initial velocity distribution and the initial spatial distribution whereas the ion mirror 
(reflector) developed in 1970’s by Mamyrin et al. corrects the initial energy distribution (Mamyrin et 
al. 1973; Mamyrin et al. 1979). 
Delayed extraction 
Delayed extraction (DE) reduces the kinetic energy distribution among ions with the same m/z ratio 
leaving the source by introducing a time lag between ion formation and ion extraction from the source, 
and at the same time it improves the pulsed beam definition(t = 0). First the ions are allowed to expand 
and separate according to their initial kinetic energy into a field-free region in the source. For ions of 
the same m/z ratio those with more initial energy will move further into the source and thereby be 
27 
 
closer to the last grid in the ion source than the less energetic ions. When the extraction pulse is applied 
after a certain delay, the less energetic ions will receive more energy because they experience a higher 
electric field than the faster and more energetic ions (see Figure 2.10). Thus, the initially less energetic 
ions receive more kinetic energy and join the initially more energetic ions at the detector. 
The ions are focused at the detector by adjusting the amplitude and delay of the pulsed extraction 
(Mamyrin et al. 1973; Mamyrin et al. 1979). Delayed extraction reduces post source decay (PSD)( 
fragmentation of the ions in the flight tube) by allowing the matrix and ionised molecules to expand 
into the source region prior to acceleration of the analyte ions. The plume density is thereby decreased, 
reducing collisional activation when the ions are accelerated into the flight tube (Kaufmann et al. 
1996). 
 
 
 
 
 
 
Figure 2.10: Delayed extraction. Schematic description of a continuous extraction mode and a 
delayed pulse extraction mode in a linear TOF mass analyser. Ions of a given mass with a correct 
energy are symbolized by ○ and ions with the same mass but to low a energy are symbolized by ●. 
Delayed pulse extraction corrects the energy dispersion of the ions leaving the source with the same 
m/z ratio. Adapted from (Hoffmann et al. 2001). 
 
 
 
 
28 
 
The reflector 
The reflector consists of a series of grids and ring electrodes, creating a retarding field that acts as an 
ion mirror by deflecting the ions and sending them back into the flight tube (Figure 2.11). It corrects 
the energy distribution of ions with the same m/z ratio because ions with higher kinetic energy will 
penetrate more deeply into the reflector than ions with a lower kinetic energy but having the same m/z 
value. The result is that ions with more kinetic energy spend longer time in the reflector than ions with 
less kinetic energy and is therefore overtaken by the slower moving ions. When the ions leave the 
reflector, the kinetic energy is restored to the same absolute value as before entering and the fast ions 
will catch up with the slow-moving ones, with the same m/z, at the detector (see Figure 2.11). 
(Mamyrin et al. 1973; Mamyrin et al. 1979). The optimal resolution is then achieved by placing the 
detector were the two ions catch up. 
 
 
 
 
 
 
 
Figure 2.11: The reflector. Schematic description of a TOF instrument equipped with a 
reflectron; (■) ions of a given mass with correct energy; (□) ions of the same mass but with a 
kinetic energy that is too low. The slower ions reach the reflectron later, but spend less time 
inside the reflector and overtake the faster ions (■). With properly chosen voltages, path lengths 
and fields, both kinds of ions reach the detector simultaneously. 
 
-3000 V
Source
Detector
-3000 V
X
Y
-3000 V +130 Vd/2 D
 
29 
 
Thus the reflector compensates for kinetic energy distribution of the ions whereas delayed extraction 
compensates for the initial time-, velocity- and spatial distribution of the ions created under the 
desorption process. The effect of delayed extraction and the reflector on mass resolution is illustrated in 
Figure 2.12. 
MALDI TOF instruments have due to delayed extraction and reflectron been able to achieve a much 
higher resolution (up to 20.000) (Mann and Talbo, 1996) resulting in an instrument, which has a good 
combination of resolution, sensitivity and fast response. One of the reasons that we are able to identify 
proteins based on Peptide Mass Fingerprinting (PMF, see later) today, is the introduction of delayed 
extraction and reflectons, which give a high mass accuracy. This lowers the level of false positives 
obtained by searching the data in the fast growing public protein sequence databases. (Jensen, et al., 
1996). 
 
 
 
 
 
 
 
 
 
 
Figure 2.12: Illustration of the improvement in resolution when the reflector and delayed 
extraction are used. The spectra were recorded on the 4700 proteomics analyzer from Applied 
Biosystems and recorded using continuous extraction, linear mode (+delayed extraction), 
 
30 
 
reflector mode (-delayed extraction) and reflector mode using both delayed extraction and the 
reflector mode. 
 
Ion Trap 
Paul and Steinwegen, who also invented the quadrupole in the 1950's, were the first to describe the 
principle of the ion trap (Paul and Steinwegen, 1960) as a slight modification of the quadrupole 
instrument. Both analyzers are based on ion motions in RF electric fields. The classical 3D ion trap, or 
Paul trap, consists of a circular electrode with two ellipsoid end caps (Figure 2.13). 
In principal an ion trap is a quadrupole in which its four rods have been bent around itself forming a 
closed loop. The inner rod forms the point at the center of the ion trap and the outer rod forms the 
circular electrode, whereas the two remaining rods form the two end caps. 
 
 
 
 
 
 
 
 
Figure 2.13: The composition of the ion trap mass analyzer. 
The ions produced in the ion source enter the trap through a small orifice in one of the endcaps (Figure 
2.14). An oscillating potential, 0, (also referred to as the fundamental RF, the sum of a direct and a 
alternative potential, see section on quadrupole) is applied to the endcaps and -0 is applied to the ring 
Ring electrode
End capEnd cap
 
31 
 
electrode. However, often 0 is only applied to the ring electrode. The potential creates a three-
dimensional quadrupolar field in which the ions are trapped (Hoffmann and Stroobant, 2001). The 
difference between the quadrupole and the ion trap is, that in the quadrupole only ions of a given m/z 
range are allowed to pass at a given time due to U and V conditions making the trajectory of these ions 
stable (see section on quadrupole). In the ion trap however, all ions of different m/z ratios are present 
and ions of a given m/z ratio are expelled from the trap at a given time by making the trajectory of these 
ions unstable (Glish and Vachet, 2003). The other ions are trapped within the system of the three 
electrodes (Figure 2.14A and B). This is due to the application of a resonant frequency along the z 
axis. Inside a quadrupole these ions would discharge themselves against the rods and be destroyed, but 
in an ion trap they repel each other, resulting in an expansion in their trajectories. The addition of 
helium gas molecules (10-3 Torr) in the ion trap absorbs any excess energy from the ions through 
collisional absorption, preventing ion losses. As with the quadrupole, the volume in which ions of a 
given m/z have stable trajectories, can be described mathematically. By sequentially making ion 
trajectories become unstable, ions are ejected from low m/z to high m/z through a small orifice in the 
opposite endcap where they are detected and used to reconstruct a mass spectrum (mass selective 
instability) (Glish, et al., 2003). 
 
 
 
 
 
 
 
 
Figure 2.14: The principle of the ion trap. The voltages applied to the system are depicted at 
C, but are present at all four times (A-D). A. Ions are trapped inside the three dimensional 
A B
DC
Scan out To 
detector
Top
Ring
Bottom
Ions in
Ions out
Trapped Selected ion
Ejected
Fragmentation
detector
electrodeelectrode
electrode
ions
ions
To 
product ion
V

detector
To 
 
32 
 
quadruple. B. An ion of a particular m/z ratio is selected. C-D. Additional steps in ion trap 
tandem ms (MS2). C. The selected ion is fragmented. D. The fragments are ejected from the ion 
trap. 
Ions can also be ejected from the ion trap by resonance ejection, where the use of higher excitation 
amplitudes increases the oscillation of a given ion resulting in its rejection from the ion trap. The 
combination of mass selective instability and resonance ejection enhances the resolution of the 
instrument (Goeringer, et al., 1992). 
Linear ion trap 
An alternative to the conventional 3D ion trap (Paul trap) is the axially-ejecting linear ion trap. The 
linear ion trap is operated as the Paul trap however this instrument is incorporated into a triple-
quadrupole instrument providing several advantages compared to the Paul trap. In the linear ion trap, a 
RF field applied to elongated electrodes is responsible for trapping the ions in the radial direction, 
whereas a static electric field on the ring-shaped electrodes at the two ends is responsible for trapping 
the ions in the axial direction. Due to a trapping stiffness in the radial direction, which is higher than in 
the axial direction, the ions will arrange themselves in a linear string in the z axis (http://heart-
c704.uibk.ac.at/linear_paul_trap.html; Hager, 2002). This improves the trapping ability of the ion trap. 
In addition, the linear ion trap is also larger than the regular ion trap providing a higher ion capacity. 
The selection of ions for fragmentation is performed by the use of mass-selective instability in 
combination with resonance ejection as described for tandem ion traps (see below). As the Paul trap, it 
can perform MSn due to the ability to store ions, while fragmenting others. In addition, the ability to 
operate as a quadrupole mass filter enables the linear ion trap to form hybrid instruments. 
One disadvantage of the iontraps is the so-called 28% rule. An iontrap is not readily able to select and 
fragment an ion and then scan out all the fragment ions. Due to the mass selective instability an iontrap 
is only able to efficiently scan out the masses beginning from 1/3 of the parant mass ion (28% rule). 
 
 
33 
 
2.4 Hybride and specialized MS instruments in proteomics 
QqTOF mass spectrometer 
In 1996 Morris and co-workers described the first Q-TOF tandem mass spectrometer. The construct 
was built by Morris to improve transmission and the efficiency to detect daughter ions, as well as to 
increase the signal-to-noise ratio. Furthermore, the aim of this construction was to improve resolution 
and mass accuracy, especially of fragment ions, in comparison with already known tandem mass 
spectrometers such as the triple quadrupole instruments (Morris, et al., 1996). They used a quadrupole 
coupled to an orthogonal-acceleration TOF instrument. The orthogonal injection was introduced in 
order to combine a continuous ion source (such as ESI) with a TOF instrument (Figure 2.15). 
O´Halloran and co-workers were the first to report of such an instrument (O´Halloran, et al., 1964). In 
orthogonal TOF, ions formed in the continuous ion source are injected along the y axis. In MS mode 
the ions travel through three quadrupoles operated in RF only mode in order to function as ion guides 
(Q0, Q1 and Q2). The first quadrupole (Q0) is an ion guide, where the ions gently collide with neutral 
gas molecules (collisional cooling or damping usually with at a pressure of 10 mTorr). This decelerates 
and focuses the ions by reducing the energy spread and the ion beam diameter giving a better 
transmission into the second and the third quadrupoles (Q1 and Q2) as well as the TOF analyzer. Q1 
and Q2 focus the ions further before they enter a field-free pusher region between a flat plate and a 
grid. This region is field-free, letting ions continue to move in their original direction in the gap. A 
voltage is then applied to the flat plate (a pusher) accelerating and thereby sending off an ion packet 
nearly in the orthogonal z direction into the flight tube. The ion packet is accelerated to several 
kiloelectronvolts per charge by a uniform DC electric field. 
The deceleration of the ions in the collision cell ensures that all ions of different m/z ratio reach the 
detector, and the pusher ensures that the ions reach the detector in an m/z dependent manner 
(Chernushevich, et al., 1999; Chernushevich, et al., 2001). Q-TOF instruments are able to isolate a 
single isotop for fragmentation, which reduces the complexity of the fragmentation spectra, when 
working with complex samples. 
 
34 
 
 
 
 
 
 
 
 
 
 
Figure 2.15: Schematic diagram of the tandem QqTOF. In MS mode the Q1 quadrupole is 
operated in RF mode only. The figure was adapted from Chernushevich, et al., 2001. 
LTQ Orbitrap XL mass spectrometer 
The orbitrap mass analyzer is with its commercial introduction in 2005 the newest type MS instrument 
in proteomic research (Hu, Q.Z., et al., Journal of Mass Spectrometry, 2005). It has a mass accuracy in 
the low- to sub-ppm range and an extremely high resolution – both comparable to the FT-ICR, which, 
like the orbitrap, also employs Fourier transformation for the calculation of ion m/z values. 
The LTQ-Orbitrap is cheaper than the LTQ-FT-ICR instrument. The running costs of the instrument 
are furthermore much lower due to the lack of the FT-ICR’s super-cooled magnet, making the 
acquisition and maintenance of the instrument in a proteomics lab much more feasible. The LTQ-
Orbitrap XL consists of a front part, which is basically a regular ESI-coupled linear trap quadropole 
(LTQ) and a back part including a so-called C-trap connected to an octopole (for higher energy 
collision induced dissociation – HCD) and the orbitrap (Figure 2.16a). 
Due to the high sensitivity, speed and MSn capabilities of the LTQ, it is the ideal partner for the 
orbitrap mass analyzer in the LTQ-Orbitrap XL instrument. The analytes are ionized in the ESI source 
Z
X
Y
 
35 
 
and guided to the LTQ through the ion optics. After manipulation in the LTQ, the ions are delivered to 
the C-trap. This is a C-shaped quadropole operating in RF mode only which functions to accumulate 
and store ions which are collisionally damped by a low nitrogen pressure before injection into the 
orbitrap by high voltage pulses (Makarov, A., et al., Anal Chem, 2006. 78(7): p. 2113-20.). 
The orbitrap is basically an ion trap, though the ions are not trapped by a radio frequency field electric 
like in a 3D ion trap or by a magnetic field as in a FT-IC. In this type of mass analyzer, the ions are 
attracted towards a central electrode (Figure 2.16a) using an electric field, an attraction 
counterbalanced by the centrifugal force arising from the initial tangential velocity of the ions 
originating from the ion injection into the orbitrap (Scigelova, M. and A. Makarov, Proteomics, 2006). 
The ions are furthermore confined by the two halves of the outer orbitrap electrode (Figure 2.16b). The 
forces imposed on the ions cause them to oscillate in complex spiral patterns not only around the inner 
electrode alone but also back and fourth in the axial direction (the z axis in Figure 2.16b). These axial 
oscillation frequencies are inversely proportional to the square root of the m/z ratios of the ions and are 
detected as an image current on the two halves of the outer electrode separated by an insulating ceramic 
ring (Figure 2.16b-c). Hereby, a single sine wave is generated by each ionic species in the orbitrap. By 
performing a Fourier transformation on the verycomplex waveforms originating from the different 
ions in the trap, the oscillation frequencies and thereby the m/z ratios can be extracted. The resolution 
of the spectra is dependent on the time of analysis, but a resolution of more than 100.000 can be readily 
achieved. 
The XL version of the LTQ-Orbitrap moreover contains a dedicated octopole collision cell coupled to 
the back-end of the C-trap (Figure 2.16a), enabling execution of high-energy collision induced 
dissociation (HCD). Hereby more extensive fragmentation than in regular, low-energy CID can be 
produced, also leading to other types of fragment ions (Olsen, J.V., et al., Nature Methods, 2007; Bean, 
M.F., et al., Anal Chem, 1991). Fragmentation of peptide ions using HCD is advantageous as the LTQ 
is incapable of efficiently trapping low-mass product ions as these cannot be stabilized in the trap – a 
phenomenon known as the “1/3 rule” indicating the loss of product ions with a mass below 1/3 of the 
precursor ion. HCD therefore facilitates the observation of the immonium ions of m/z 216.04 
indicating a phosphotyrosine or the reporter ions from the iTRAQ® reagent used for labeling of 
peptides in quantitative proteomic experiments (m/z 114-117). The intensity of the ions detected after 
36 
 
performing HCD, is nevertheless an order of magnitude lower than by CID in the LTQ, due to 
transmission loss. In addition, the number of ions required for the HCD is significant higher than for 
normal CID in the LTQ. Thus this is still leaving room for major improvements of the application of 
the technique in analysis of low-abundant and complex samples. 
Recently, the LTQ-orbitrap XL has been equipped with capabilities for performing electron transfer 
dissociation (ETD), a gentle fragmentation technique which allow efficient fragmentation of multi-
charged peptides and proteins (see later). 
The very high mass accuracy of the instrument (<5 parts per million (ppm) with external calibration) 
can be improved even further by internal calibration. The polycyclodimethylsiloxane polymer 
(445.120025 Da), which is always present in the ambient air, and is therefore an inevitable ion showing 
up in mass spectra from ESI instruments (Schlosser, A. and R. Volkmer-Engert, Journal of Mass 
Spectrometry, 2003), has been used for internal calibration in real-time, employing its mass as a so-
called lock-mass (Olsen, J.V., et al., Molecular & Cellular Proteomics, 2005). This method relies on 
filling the C-trap with a particular amount of polycyclodimethylsiloxane ions and scanning them along 
with the analyte ions. The average mass accuracy can thus be reduced to the sub-ppm level by 
correcting all measured masses by applying the same ppm deviations for all masses as for the lock-
mass. 
 
 
 
 
 
 
 
 
 
 
 
A)
B)
Orbitrap
HCD collision cell
37 
 
Figure 2.16: A) The LTQ-Orbitrap XL consists of an ion source, ion optics, and a linear ion 
trap connected to the C-trap through the front end, and with the HCD collision cell in the back 
end. From the C-trap, the ions are injected in to the orbitrap. B) The orbitrap mass analyzer. a) 
Central electrode b) Outer end-cap electrode c) Insulating ceramic ring.. 
 
Fourier transform Ion Cyclotron Resonance mass spectrometer (FT ICR) 
In 1974 Fourier transform (FT) MS was described by Comisarow and Marshall (Comisarow and 
Marshall, 1974a and b). The FT instrument consists of four components: The first component is a 
magnet creating a uniform high magnetic field of 4.7 – 13 T, the second component is an analyzer cell, 
where ions are stored, analyzed and detected. The analyzer cell can be a cubic cell consisting of six 
plates positioned inside the magnetic field (Figure 2.17). The third component of an FT instrument is 
an ultra-high vacuum system. Ions generated in the ion source are guided through compartments with a 
series of pumps to gradually lower the pressure before the ion enters the ICR cell. At this point pressure 
will be in the range of 10-13 to 10-14 Bar, and the temperature about absolute zero. The fourth 
component is a sophisticated data system (Amster, et al., 1996). 
As the ions generated in the ion source enter the analyzer cell, they are exposed to the magnetic field 
and to an electric field generated between the electrodes of the analyzer cell. This forces the ions into a 
circular motion (the cyclotron motion) in a plane perpendicular to the field (Figure 2.17). 
The cyclotron motion arises from the interaction of an ion with an unidirectional magnetic field and is 
due to the Lorentz Force: 
F = zv x B 
Where F is the Lorentz Force observed by the ion, when it enters the magnetic field, z the charge of the 
ion, v the velocity of the ion, and B the magnetic field strength (constant). x indicates the cross product 
and not the multiplication symbol. The electric field acts to force the ions away from the center of the 
analyzer cell, whereas the magnetic field prevents the ions from colliding with the walls of the analyzer 
cell. The frequency of this magnetron motion is m/z dependent: 
38 
 
c = zB / 2m 
c being the induced cyclotron frequency, and m the mass of the ion. By inducing a swept RF pulse 
across the excitation plates a packet of ions of a given m/z ratio is excited. By adjusting the pulse to 
have the same frequency as the cyclotron frequency of the ions, ions will absorb energy. This excitation 
results in an increase in the radius of the ion motion bringing them closer to the detector plates, where 
they induce an image current between the detector plates. The frequency of the image current is in 
resonance with the cyclotron frequency of ions of a particular m/z ratio, and the intensity is 
proportional to the number of ions. Ions of one specific m/z ratio are excited together and the image 
current is recorded and analysed by the data system. When the RF goes off resonance for that particular 
m/z ratio, the ions return to their initial motion (relax) and other ions can be excited. The RF sweep can 
also be adjusted to consider all frequencies simultaneously, resulting in a measurement of all the ions at 
the same time. 
This creates a complex frequency vs. time spectrum of all the signals. Deconvolution of these signals 
by FT methods results in a deconvoluted frequency vs. intensity spectrum, which can then be converted 
to a mass spectrum by the following equation: 
m/z = B / 2c 
 
 
 
 
 
 
 
 
Magnetic
Field, B
Trapping
plates
Detector
plates
Excitation on  one frequency (RF) excites one m/z
convoluted
frequency
spectrum
frequency
deconvoluted
spectrum
Mass
spectrum
Induced alternating
currentExcitation
plates
RF
FT MC
Ion 
Analyzer cell
 
39 
 
Figure 2.17. The principle of FT ICR mass spectrometry. 
 
The FT ICR is a powerful instrument with many possibilities. One great advantage of FT MS is the 
ion-trap nature of the instrument, which allows it to measure ions without destroying them. This makes 
it possible to perform subsequent fragmentation or perform other studies on the ions. Another 
dimension to the use of FT ICR mass spectrometry was the introduction of Electron Capture 
Dissociation (ECD), where charge-mediated fragmentation is produced. For proteins, this 
fragmentation occurs solely in the backbone, sparing the side chains, which make it an efficient method 
for the site-specific analyses of labile posttranslational modifications that are usually lost by traditional 
fragmentation (Sze et al., 2002). Proteins containing cysteine residues are an exception, due to their 
high radical affinity. ECD also enables the MSn analysis ofentire proteins and their larger fragments 
(“the top down” approach) as apposed to the traditional analysis of low molecular weight proteolysis 
products (“the bottom up” approach) (McLafferty et al., 2001) 
The FT instrument can also be combined with electrospray ionization giving high throughput 
measurements and also the possibility of coupling with separation techniques such as high-performance 
liquid chromatography (HPLC) (Schmid, et al., 2000). 
 
Linear ion trap- Fourier Transform (LTQ-FT) 
Prior to the development of the LTQ-orbitrap the most sophisticated and accurate MS instrument for 
performing protein identification ad quantitation was the LTQ-FT instrument which shares some of the 
characteristic and features as the LTQ-orbitrap. The difference is the speed of the analysis and the cost 
as the orbitrap do not need to be cooled by liquid helium which is a significant cost for the FT 
instruments. An illustration of the LTQ-FT instrument is shown in Figure 2.17. 
 
 
 
40 
 
 
 
 
 
 
 
 
 
 
Figure 2.17. The construct of the Finnigan LTQ-FT instrument. 
 
2.5 Mass spectrometric sequencing of peptides – tandem MS (MS/MS) 
MS driven proteomics is based on the ability to measure the molecular weight of proteins and peptides 
and to select these for fragmentation by MS/MS to obtain detailed sequence information. In order to 
retrieve structural information on peptides/proteins using MS, the peptide/protein needs to be 
fragmented into smaller parts that reflect the structure and modifications of the original peptide/protein. 
The m/z ratio of these fragments can then be compared to known m/z ratio of particular chemical 
structures stored in databases. This can most readily be achieved if the sequence of the peptide/protein 
is known prior to the analysis from for example genome sequencing. If the sequence of the protein is 
not known one can, based on the sequence information obtained from the tandem MS sequencing, 
search for homologe protein sequences in a database (using e.g., BLAST search), or generate enough 
sequence for designing primers for sequencing the particular protein – a procedure which is becoming 
very fast and in-expenseable. 
210L/sec 210L/sec60 m3/hr
Triple Ported Turbo
400L/sec300L/sec25L/sec
x
y
z
7 T Actively Shielded
Superconducting
Magnet
ICR Cell
2D Ion Trap
 
41 
 
 Fragmentation of peptides/proteins is achieved by transferring extra energy to the stable ions formed 
during ionization (see below). Extensive fragmentation can also be induced during ionization by the 
use of highly energetic ionization techniques. There are many different ways to nduce fragmentation of 
the peptide, e.g., collision induced dissociation/collision activated dissociation (CID/CAD), post source 
decay (PSD), Electron transfer dissociation (ETD), etc. Below is described the basic nomenclature of 
peptide fragmentation and the basis for some of these fragmentation techniques. 
 
Nomenclature for peptide fragmentation 
In 1984 Roepstorff and Fohlman proposed a common nomenclature for the different fragments 
obtained (Roepstorff and Fohlman, 1984). This nomenclature was modified in 1988 by Biemann 
(Biemann, et al., 1988). The cleavage of a bond in the main chain can occur at three different positions, 
in a C-C bond, a C-N bond or in an N-C bond. The resulting daughter ions can be divided into two 
separate groups. Those containing the N-terminal of the original peptide and those containing the C-
terminal of the original peptide. These two groups can each be further divided into 3 subgroups. The 
group of N-terminal containing fragments includes an, bn and cn ions depending on what position on the 
peptide backbone fragmentation has occurred. The group of C-terminal containing fragments includes 
xn, yn and zn ions. n indicating the number of amino acids in the fragment. An overview of the different 
fragment ion that can be obtained by MS/MS of peptides is showed in Figure 2.18. 
 
 
 
 
 
 
 
H2N C
R
H
+
A. Peptide fragmentation
B. Immonium ion
H2N CH
R1
C
O
NH CH C
R2 O
NH CH C
R3 O
OH
a1 b1 c1 a2 b2 c2
x2 y2 z2 x1 y1 z1
H+
H2N C
R
H
+
A. Peptide fragmentation
B. Immonium ion
H2N CH
R1
C
O
NH CH C
R2 O
NH CH C
R3 O
OH
a1 b1 c1 a2 b2 c2
x2 y2 z2 x1 y1 z1
H2N CH
R1
C
O
NH CH C
R2 O
NH CH C
R3 O
OH
a1 b1 c1 a2 b2 c2
x2 y2 z2 x1 y1 z1
H+
42 
 
Figure 2.18: Common nomenclature of peptide fragment ions. The common nomenclature 
for different fragment ions generated in MS/MS was proposed by Roepstorff and Fohlman in 
1984, and slightly modified four years later by Biemann. In CID, primarily y and b ions are 
produced, whereas z and c ions are generated by ECD and ETD. 
There are two additional classes of peptide fragments; fragments created by the cleavage of two bonds 
in the peptide chain, and fragments that are further fragmented in the amino acid lateral chain. 
Fragments of the first class are referred to as internal fragments, since they have lost both the N- and C-
terminal sides. They are usually no more than three or four amino residues in length and they are 
especially abundant if the amino acid proline is present in the peptide sequence. Internal fragments are 
designated with series of simple letters representing the sequences of the fragments. Fragments of the 
second class constitute the immonium ions of individual amino acid residues (Figure 2.18). They 
appear in the lower mass region and are not equally abundant for each amino acid residue, but some are 
commonly seen and these provide important information about a sequence. The immonium ion from 
the tyrosine residue at m/z 136 is one of the immonium ions commonly observed and this ion is 
important, when looking for phosphorylated tyrosine residues in a sequence. Immonium ions are 
represented by the capital one-letter code of the corresponding amino acid residues. Immonium ions are 
not frequently observed in iontrap instruments due to the 28% rule. Three additional fragment ion 
types can be observed in high energy spectra, dn, wn and vn. dn and wn and they are formed by the 
cleavage of the bond between the  and the  carbon atoms of the amino acid side chains, dn having the 
positive charge on the N-terminal and wn having the positive charge on the C-terminal fragment. The vn 
fragment ions are formed by the loss of the entire side chain. 
 
Collision induced dissociation (CID) 
Collisional induced dissociation (CID), also known as collisional activation dissociation (CAD), is 
accomplished by converting some of the kinetic energy from the peptide into vibrational energy by 
collision of the (M + nH)n+ ion with a neutral gas (e.g. Ar, Xe or He) in a collision cell (Shukla and 
Futrell, 2000)(Figure 2.19). The fragmented ions, termed daughter ions, are then separated in a second 
mass analyzer and detected by the detector, giving raise to a fragment ion spectrum. The number and 
43 
 
nature of the ions detected will depend on the collisional energy used in the collision cell, on the size of 
the peptide and on the amino acid sequence of the peptide. High energy collision induced 
fragmentation (several keV), as used by TOF-TOF (Medzihradszky, et al., 2000) and magnetic sector 
instruments (Hoffmann, 2001), induce extensive fragmentation and give rise to other types of 
fragments ions. These hold a lot of information, but also complicate the assignment of spectra to amino 
acid sequences. Low collisional energy ( 100 eV) as used by quadrupole and ion trap instruments, 
induce less fragmentation and subsequently