13 Chapter one: Recent development in protein affinity selection mass spectrometry
13
Chapter one:
Recent development in protein affinity selection mass
spectrometry
Chapter one
14
Abstract
This review intends to provide an overview of protein affinity selection methodologies
that can be coupled to mass spectrometry. It will shortly address the principle of each of
the methodologies, but will mainly focus on the developments in recent years, and the
applicability of the methodologies to lead generation or other research fields important in
drug discovery.
Introduction
The development of new lead compounds in drug discovery has been of
continuous importance over the past decades. However, high attrition rates and the
decreasing number of new drug approvals in the past years have increased the necessity
for new development tools. Protein affinity selection methods utilizing mass
spectrometry are amongst the more recently developed methodologies that can be a
valuable addition to traditional drug discovery techniques. They distinguish themselves
by utilizing the very high sensitivity and selectivity that is inherent to mass spectrometric
detection, while retaining the biological specificity that is typical for commonly used
plate reader assays.
The use of mass spectrometry has a number of consequences for the bio-assay. A
positive implication is the fact that no labels are required in mass spectrometry. This
widens the application area for these methods, including target proteins for which no
label is available or can be developed. A second advantage is that the mass spectrometer
measures the actual compound that has the affinity for the protein, instead of a labeled
competitor. Since protein selection methodologies deliver simultaneous biological and
structural data on the compound to be analyzed, the detection of false positives due to
library impurities or degradation of the compounds in the mixture is very improbable.
Several methodologies have been developed using direct measurement of a
protein-ligand complex in mass spectrometry, and a number of interesting reviews have
been published recently.[1-3] However, many of these methods require extensive
Chapter one
15
optimization in order to combine protein-ligand complex stability with conditions
suitable for mass spectrometric detection. Secondly, advanced mass spectrometric
method development is required to distinguish the protein-ligand complex from the
unbound protein without prior separation. As a result, these methodologies have not yet
shown to be universally applicable to a large number of target proteins. However, they do
provide the most direct evidence of protein-ligand complexation, provided the
assumption that gas-phase complexation in the mass spectrometer can be used as a model
for in vivo complexation.[4] Furthermore, many protein-ligand binding assays do not use
mass spectrometry. An excellent review on this topic, published by de Jong et al., focuses
on assay technology used in high throughput screening, with a strong emphasis on
photometric detection.[5]
In this paper a comprehensive overview of recent developments in the field of
protein affinity selection methods that utilize mass spectrometry is provided. The scope is
limited to methodologies that involve a step that separates bound and unbound protein
before detection. All these methodologies consist of the same four steps: (1)
Complexation of the protein and test ligand. (2) Separation of non-bound compounds
from the protein-ligand mixture. (3) Elution/dissociation, resulting in the release of the
free, bioactive ligand, and (4) Detection of the eluted ligand by mass spectrometry. An
overview is provided of the various affinity selection methodologies that are being
applied in modern-day drug discovery, such as size-exclusion chromatography, (pulsed)
ultrafiltration, surface plasmon resonance and immobilized or dynamic protein affinity
selection methodologies.
Affinity Chromatography
Affinity chromatography is a process in which a compound, be it a protein, small
molecule, antigen, antibody or any such biologically active agent is immobilized on a
solid support. When a mixture of analytes or a complex sample containing an analyte is
injected onto the column containing this solid support, the analyte that shows affinity for
the immobilized compound gets retained, and separated from the rest of the sample. The
Chapter one
16
method originates from the late sixties, when it was used to extract and purify
enzymes[6] or antibodies[7]. These methodologies were based upon solid supports
consisting of mainly sepharose or agarose. These materials were known for their low
non-specific adsorptive properties, and their relative ease of modification, allowing easy
immobilization of the desired target. However, when the methodology developed towards
affinity chromatography, with the aim of not only extracting and purifying the analytes,
but also ranking them according to their affinity, it became a necessity to use support
materials that were better equipped for the flow rates and capillary pressure that is
common in HPLC. The use of pressure resistant solid supports is often referred to as the
defining advantage of high performance affinity chromatography over conventional
affinity chromatography, and was a starting point for a rapid expansion of the number of
applications published in this field.[8]
Many modern-day innovations are aimed at improving the immobilization of the
target on the backbone and improving the quality of the materials used in the solid
support. However, most of them use two approaches for the actual proof of principle. The
first is frontal affinity chromatography, in which a known concentration of analyte is
infused onto an immobilized protein column, and affinity is calculated based on the
saturation time and the shape of the breakthrough curve. The second is zonal elution, in
which a small amount of analyte is injected onto an immobilized protein column together
with a known amount of a competitor. By varying the concentration of the competitor
and measuring the retention time of the analyte, its affinity can be calculated.
Frontal affinity chromatography
The concept of frontal affinity chromatography is more complicated than it
appears at first sight. A target, very often a protein or receptor, is covalently immobilized
on a column. An analyte is then infused onto the column at a known concentration, and
the analyte concentration exiting the column is measured. In the beginning this amount
will be very low, because there is a large amount of free receptor for the analyte to bind
to. However, as a larger fraction of the immobilized receptor is bound, the amount of
analyte being measured will slowly increase. At some point, all the receptor on the
Chapter one
17
column will be bound to the analyte, and the signal in the detector will equal that of the
concentration of analyte that is being infused. However, the speed at which this happens
is dependant on the association (Ka) and dissociation constants (Kd) of the protein-analyte
interaction. An analyte with fast dissociation kinetics will have a longer saturation time
than one with slow dissociation, because during the process some of the protein-analyte
complexes will dissociate, increasing the fraction of unbound protein. As a result the
affinity of the analyte for the protein can be calculated by varying the concentration of
analyte, and determining the absolute amount of analyte needed to reach 50% saturation
for every concentration. A typical result for such an experiment is shown in figure 1.1[9].
Figure 1.1: A frontal analysis experiment performed with L-tryptophan on an immobilized human
serum albumin column. The concentration of applied L-tryptophan (from left to right)
was 100, 50, 25 and 12.5 µM. The flowrate was 0.25 mL/min and t0 was 3.6 min.
Reproduced from ref [9].
When developing a FAC method, the handbook of affinity chromatography by
Hage et al[8], and the paper on practical protocol development by Ng et al.[10] should
provide all the theoretical and practical background required. This review will focus on
the most significant developments in FAC methodologies that employ mass spectrometric
detection. These developments are currently focused on two fundamental assay
properties: the improvement of the solid support material to facilitate easier protein
Chapter one
18
immobilization, reduce non-specific binding and stabilize certain classes of target
proteins, and the miniaturization of the protein affinity column, in order to minimize
protein consumption. Prime examples are the use of artificial immobilized membranes
(AIM), monolithic affinity columns, and miniaturization of the affinity columns. Next to
these fundamental improvements, a significant number of papers have been published
expanding the number and types of targets successfully immobilized.
The most significant development in the field of novel solid supports has been the
application of immobilized artificial membranes (IAM) to form immobilized receptor
columns. Ever since the introduction of FAC in conventional HPLC systems, the target
proteins were immobilized mainly on silica, glass or polystyrene material. One of the
largest disadvantages of these solid support materials was the fact that many proteins lost
a major part of their activity upon immobilization. In order to widen the applicability of
FAC, a solid support contributing to protein stability was required. IAMs were developed
by Pidgeon et al. in 1989 [11], consisting of silica beads covered with membrane lipids to
be used as a stationary phase in chromatography. Many receptor types have been
immobilized on IAMs since then, and in recent years Beigi et al. have shown the
immobilization and screening of analytes to G-protein coupled receptors.[12,13].
Recently, Temporini et al. have successfully immobilized the orphan receptor GPR17 on
artificial immobilized membranes, including an affinity ranking experiment, the results of
which are shown in figure 1.2.[14] Also, Moaddel et al. published a comprehensive guide
on IAM development and preparation, allowing fast implementation by groups with little
experience in the field.[15]
The other two major developments are often applied in conjunction. Because
columns on microfluidic chips or in nano-LC scale experiments are very small, they have
very low protein consumption. Furthermore, because these small columns are hard to
pack with conventional bead type materials, they have been a prominent target for the
development of monolithic affinity columns. A very elegant method requiring complex
implementation was published by Besanger et al. immobilizing membrane-bound
nicotinic acetylcholine receptor in a monolithic column built up from diglycerylsilane
polymers.[16] By adding the right amount of polyethylene glycol, the membrane and the
active receptor were wholly incorporated in a silica monolith, and successfully used in a
Chapter one
19
FAC-MS experiment comparing the affinity of epibatidine to a benchmark method.
However, slight adjustments in the manufacturing procedure could lead to major changes
in the monolith formation and the receptor activity. Okanda et al. published a monolithic
silica column with immobilized lectins incorporated with suitable properties for affinity
ranking in both nano-LC and capillary electrophoresis.[17] Finally Feng et al. reported
the fabrication of a monolithic silica column with incorporated Fe3+
, allowing nano-scale
immobilized metal affinity chromatography (IMAC).[18]
Figure 1.2: Affinity ranking of 3 ligands to GPR17 receptor immobilized on an IAM using frontal
affinity chromatography-mass spectrometry. From top to bottom UDP (Kd = 1140 nM),
MRS2197 (Kd = 508 nM) and cangrelor (Kd = 0.7 nM). Reproduced from ref [14].
Zonal Elution
Zonal elution is the most straightforward method of performing affinity
chromatography. Basically it is a classic competition experiment, implemented in a
HPLC system. A known and constant amount of the analyte is injected in a small amount
onto the affinity column, in the addition of a competitor. When a series of experiments is
performed using a varying concentration of competitor, the retention of the analyte will
vary. For a high concentration of competitor, analyte retention will be low, because a
Chapter one
20
large fraction of the available protein bound to the column is occupied by the competitor.
Likewise, for a low concentration of competitor, analyte retention will be high. The
advantage of this method over FAC is the small amount of analyte consumption, which is
useful when analytes are scarce, expensive or hard to synthesize. However, frontal
analysis provides more information per experiment, allowing for instance the
determination of the number of binding sites per column and the association constants of
the protein-analyte complexes. Because of the limited amount of data collected, most
affinity chromatography development in pharmaceutical industry is aimed at frontal
analysis. Zonal elution is nowadays more often used as a means to acquire additional data
on an interaction observed in frontal analysis, such as the location of a binding
site.[19,20] Furthermore, many of these methodologies involve other means of detection
than mass spectrometry. One of the few recently published method based solely on zonal
elution has been developed by Bertucci et al., who investigated the binding dynamics of
HIV inhibitors to human serum albumin using zonal elution.[21].
In conclusion, it is apparent that frontal affinity chromatography is a prominent
technique in pharmaceutical research, because of the high amount of data produced, and
the wide variety of targets that can be immobilized. In recent years, FAC applicability has
extended to include G-protein coupled receptors, a hot target in pharmaceutical research
and notoriously hard to stabilize when used in bio-assays. Due to the fast development of
frontal analysis, zonal elution is used more as an auxiliary technique. When comparing
frontal analysis to other affinity selection techniques, two drawbacks of the technology
must be addressed. First, for every target that is immobilized, extended controls must be
performed to check whether the immobilized protein has the same functionality as the
protein in native conditions. Immobilization influences the freedom of movement of the
protein, and might impede binding to certain binding sites, thus decreasing the affinity of
certain ligands to the protein, causing false negatives. Secondly, immobilized protein
columns have a very limited stability. If an HTS screening assay would be developed
using FAC, it would require a large number of functionally equal columns to be
produced, which might be challenging.
Chapter one
21
Affinity Selection
Affinity selection, also known as affinity capture or affinity trapping
methodologies, are similar to affinity chromatography in the sense that their mechanism
of action depends on possible analytes binding to immobilized proteins on a solid
support. However, in affinity selection no chromatographic separation of ligands is
involved. The conditions in which the binding occurs are optimized to stabilize the
protein-analyte complex, wash off non-binders, and then dissociate the complex to detect
the binding analytes. In order to identify binders, most of these methodologies employ
mass spectrometric detection. Because the compounds are not separated based on their
affinity for the receptor, the resulting analytes can not be ranked, but will only be
grouped in binders and non-binders. Optimizing the threshold to distinguish these two
groups, in order to eliminate false negatives, and limit false positives, is one of the most
challenging aspects of affinity selection development.
Although affinity selection in one form or another has been around for decades
[22-24], it has become more prominent in bio-analysis upon the introduction of the lectin-
based affinity materials, used to extract bacteria and carbohydrates with a specific
binding motif for MALDI-TOF analysis.[25,26]. An example of results obtained using
these lectin-based affinity capture surfaces is shown in figure 1.3. These methodologies
allowed extremely sensitive detection of target proteins that were hard to analyze in any
conventional way due to the complex matrices they were in. Since that time a number of
very specific affinity materials have been developed and employed with considerable
success. Krugman et al. presented their work in which a novel family of proteins found in
leukocytes bound specifically to phosphoinositide affinity material, and could be isolated
and identified for the first time.[27] Kong at al. described a diamond based affinity
material with the property of extracting any kind of protein from any complex mixture.
It’s lack of specificity was countered by a direct coupling to a MALDI-TOF, allowing
mass spectrometric identification.[28] Recently, Ferrance has shown the feasibility of
immobilizing proteins on gellan beads (a polysaccharide).[29] The optical transparence
of these beads allows direct measurement of the amount of protein successfully
Chapter one
22
immobilized on the material. Freije et al. developed a sepharose material with a small
peptide immobilized on it in order to select metalloproteases based on their activity [30],
and successfully implemented this material as a sample enrichment step in an online
affinity selection-trypsin digestion-MS proteomics approach able to identify the proteases
and rank their activity [31]. Finally, Johnson et al. utilized affinity selection by
immobilizing an in-house developed capture ligand to selectively extract protein A from
Staphylococcus Aureus, resulting in an easy to optimize affinity material synthesis
procedure that can be used on a wide variety of target proteins.[32]
Figure 1.3: MALDI-TOF mass spectra resulting after the extraction of Sinbis virus (MW = 51800)
from cell culture using (a) lectin-modified affinity capture surface, and (b) unmodified
affinity membrane. BSA is bovine serum albumin. Reproduced from ref [25].
Chapter one
23
Immobilized Metal Affinity Chromatography (IMAC) comprises a large number of
methods similar to protein affinity chromatography, but with one difference putting it
beyond the scope of this paper. It separates proteins based on their affinity to an
immobilized metal ion on a solid support. Instead of protein ligand affinity, metal-ligand
affinity is determined. This separation mainly depends on the number and location of
histidine residues in the proteins to be analyzed, and to a lesser extent cysteine and
tryptophane residues.[33] However, when a protein is His-tagged, a chain of six
histidines is artificially implanted into the protein, resulting in a very high affinity
between the His-tag and the immobilized metal ion. Because of this high affinity, there is
no longer any chromatographic separation involved, the IMAC column simply traps any
His-tagged protein from any matrix. As a consequence, nowadays this is the most widely
used method of protein purification. A perfect recent example of which is shown by
Cheek et al. who developed a monolithic IMAC column in order to purify His-tagged
lentiviral vectors.[34] Jonker et al. developed a dynamic protein affinity selection
methodology utilizing immobilized metal ions, by trapping an in-solution formed protein-
ligand complex on a small nickel-loaded column.[35] This allows complex formation
under native conditions, using the immobilized metal column merely to separate the
bound and unbound fraction, followed by mass spectrometric detection.
Another variant of affinity capture methodologies that has been increasingly
popular is ligand fishing using magnetic particles. A target protein is immobilized on
magnetic beads, and added to an oftentimes complex mixture of possible ligands. All
possible ligands are allowed to bind to the immobilized protein, and the mixture is
exposed to a magnetic field, retaining the beads and anything bound to them, while the
unbound fraction is removed. The regular use of magnetic beads for ligand fishing
originates in the early nineties, when streptavidin-coated beads were first used to purify
biotin-labeled proteins[36], and antigens immobilized on magnetic beads were applied
for antibody purification.[37]. Within a couple of years this lead to two major
developments: immobilized metal magnetic beads, able to trap His-tagged proteins[38],
and the paper by Mehta et al. first describing direct immobilization of proteins on
magnetic beads.[39] Nowadays, most of these types of magnetic particles are being
produced for the mass-market, and are available to anyone. This has lead to a rapid
Chapter one
24
increase in the number of papers published utilizing magnetic beads. One example of a
recent development of the use of immobilized metal magnetic particles is magnetic bead
dynamic protein-affinity selection, in which the beads are used to trap a His-tagged
protein-ligand complex from a sample, allowing protein affinity selection in much more
complex mixtures, the results of which are shown in figure 1.4.[40]
Figure 1.4: Weak binder identification using magnetic bead dynamic affinity selection as sample pre-
treatment for a mixture of possible ligands. Norethisteron is successfully extracted as the
only binder in the set of compounds. Reproduced from ref [40]
Furthermore, Meyer et al. developed a gradient magnetic fishing methodology utilizing a
non-linear magnetic field to capture superoxide dismutase from crude whey.[41] In the
field of protein immobilization on magnetic particles the work of Marsza et al is a prime
example of the vast body of work published by the group of Wainer et al., employing
Chapter one
25
magnetic beads coated with heat shock protein, and effectively using them for ligand
fishing.[42] Finally, Hu et al. published an enzyme inhibition screening assay utilizing
enzymes immobilized on magnetic beads with considerable success.[43]
In conclusion, affinity selection methodologies are rapidly developing, and are
important tools for future drug discovery. Their advantage is the wide applicability of
affinity selection, the easy compensation that can be made for non-specific binding, and
their relatively high sensitivity. The major drawback is the fact that affinity ranking using
these methodologies is not possible. The results of these methods will be a group of
binders and a group of non-binders. The biggest challenge is to eliminate false negatives,
and limit false positives as much as possible. In order to rank the affinities of the group of
binders, most of the other methods in this paper are more suitable.
Ultrafiltration
Traditional
In the early eighties, ultrafiltration was developed as a means to measure protein-
ligand interactions in solution based complexes.[44,45] The concept of the technique is
rather straightforward. A certain amount of pressure is applied to an amount of liquid
containing protein-ligand complexes as well as free ligand and protein. This can be
achieved by applying pressure using a pump, vacuum or centrifugal force. The
ultrafiltration unit contains a molecular weight cut-off membrane, allowing solvents and
non-bound small molecules to pass through, but retaining proteins and protein-ligand
complexes. This achieves effective separation of the bound and non-bound fractions of
analyte, allowing detection of the bound ligand by any available detection method. The
large advantage of this method over other existing methodologies is the fact that complex
formation takes place in solution, allowing the protein the same degree of freedom it
would in vivo. Most competing methods at that time used immobilized proteins, with the
drawback that immobilization could influence the binding properties of the protein.
Chapter one
26
Since that time, many interesting developments have followed. Wieboldt et al.
developed an online ultrafiltration MS methodology to identify affinity for an antibody in
a testset of structurally very similar compounds (figure 1.5).[46]
Figure 1.5: Typical results of a immunoaffinity ultrafiltration methodology. The testset consists of
20-30 benzodiazepines. In (a) anti-nitrazepam is used as the affinity ligand before
ultrafiltration. In (b) it is anti-flunitrazepam. (c) is the LC-MS reference chromatogram of
the complete testset. Reproduced from ref [46].
Weisiger et al. developed a methodology in which the complex in vivo interaction
between bilirubin and albumin is studied utilizing serial ultrafiltration.[47] In order to
study this interaction, a series of successive ultrafiltration steps is required in order to
remove impurities in the sample. This resulted in very accurate affinity constant, allowing
the study of factors such as albumin concentration and pH, which have relatively small
influence on the affinity constant. A number of groups have developed methodologies
using continuous ultrafiltration.[48,49] In this technique, a fixed amount of protein is
injected into an ultrafiltration chamber. The analyte is then pumped through the chamber.
Much like frontal affinity chromatography, at first the analyte will bind to the protein
available in the ultrafiltration unit, but upon binding the amount of available binding sites
Chapter one
27
will decrease, and an increasing amount of analyte will pass through the molecular
weight cut-off membrane and be detected. A recent example of these methods using
automated continuous ultrafiltration is published by Heinze et al.[50], utilizing it to
determine antibody-protein binding in human plasma. Fung et al. were one of the first to
automate ultrafiltration for high throughput analysis for Schering-Plough.[51] Some other
recent publications include the work of Comess et al, who developed a high throughput
serial ultrafiltration methodology to screen a library of compounds for affinity to a
pharmacologically relevant streptococcal enzyme.[52] Finally, Li et al developed an
online coupled ultrafiltration LC-MS methodology that was used to screen natural
extracts for �-glucosidase inhibitors.[53]
Pulsed Ultrafiltration
The development of pulsed ultrafiltration by van Breemen et al. significantly
increased the applicability of ultrafiltration.[54] Instead of filtration of an entire sample, a
small amount of sample was injected into the ultrafiltration unit. The liquid flow would
push the non-bound fraction through the molecular weight cut-off membrane, to waste.
Afterwards the bound ligand is dissociated from the protein by buffer adjustment. The
bound fraction is then pushed through the membrane, and immediately measured and
identified by mass spectrometry. The two main advantages over traditional ultrafiltration
are that: (i) less analyte was needed; (ii) receptors could be reused if a non-destructive
dissociation buffer was applied. The methodology could be applied to complex mixtures
as well as combinatorial libraries, and it was functioning online, automated and suitable
for high throughput screening. Since the publication of this methodology, most novel
ultrafiltration applications have used some or all of the characteristics of pulsed
ultrafiltration, very often without identifying them as such. As a result, there no longer
exists a strict division between traditional and pulsed ultrafiltration. The technology and
terminology involved are very often used incorrectly. However, a significant number of
interesting advancements has been published since.
Some successful applications of pulsed ultrafiltration include the work of Shin,
Liu and Cheng, all in the group of van Breemen, who used pulsed ultrafiltration to study
metabolic stability[55], inhibitors of protein aggregation[56] and ligands for human
Chapter one
28
retinoid X receptor alpha[57]. In the field of ultrafiltration, most development is focused
on new applications, and not on improving the technology.
When reviewing ultrafiltration, it is apparent that the two biggest advantages of
ultrafiltration based affinity selection methodologies are (i) in solution complex
formation, and (ii) the possibility to reuse the receptor. However, in practice the most
common problems occurring when implementing ultrafiltration are leakages, protein
denaturation and consequentially reduced activity of the receptor mixture and non-
specific binding to the ultrafiltration membrane. As a consequence, the use of competing
technologies is growing faster than the use of ultrafiltration.
Size exclusion Chromatography
The concept of separating compounds by size has been developed around 1964, in
the form of gel permeation chromatography, mainly applied to the analysis of high
molecular weight polymers.[58] The concept is as simple as elegant. A cross-linked
dextran gel is used as column material. The porosity of the gel will determine its
separation properties, resulting in less retention for larger molecules, and more retention
for smaller molecules. This is caused by the diffusion of the smaller molecules into pores
in the polymeric gel. Because small molecules can enter these pores and the larger
(proteins) can not, their retention will be longer than that of the protein. With the
development of more advances gels and other size exclusion materials, the separation
power of size exclusion-based methods has steadily increased since their inception.
Mainly due to favorable results obtained in drug discovery projects performed by the
major pharmaceutical corporations, size exclusion affinity selection coupled to mass
spectrometry is one of the most important protein-ligand interaction screening
methodologies in current pharmaceutical research. Size exclusion methods are easy to
automate and very suitable for implementation in High Throughput Screening (HTS)
assays. As a result, they are the fastest methods of all discussed in this paper.
Most gel permeation chromatography has been developed in order to function in
spin columns, in which the sample is pipetted onto the gel, but will not permeate the gel
Chapter one
29
unless exposed to centrifugal force.[59,60] The most direct successor to the classic gel
permeation chromatography is SpeedScreen technology developed by Novartis.[61] It
consists of a double 96-well plate format. The upper contains size exclusion gel, and has
holes in the bottom of the plate. An in-solution protein-ligand incubation mixture is
pipetted on top of the gel, and it is placed upon the second plate, a collection plate. The
incubate is separated due to centrifugal force, and bound and unbound ligands are
separated (figure 1.6). The 96-well plate is then analyzed using a standard LC-MS
system. The system screens one 96-well plate within 10 minutes, has routinely been
applied by Novartis and resulted in a number of lead compounds.[62] However, the
preparation of the 96-well plates is an intricate process, and might complicate its
application outside the laboratory where it was developed. Since its development, no new
developments have been published on the subject by Novartis or any of their competitors.
Figure 1.6: Results obtained using the Speedscreen methodology. Olomoucine and Staurosporine,
both known ligands for Protein Kinase A are isolated and detected from a mixture of 400
non-binding compounds. Reproduced from ref [61].
Chapter one
30
Based on the same concept, size exclusion columns are being used in almost all
branches of protein analysis. A much larger variety of materials is available in size
exclusion chromatography than in gel permeation chromatography (which traditionally
uses Sephadex gel), and the reusability of these materials allow the use of these columns
in online LC methods. In 2004, Neogenesis developed the first automated HTS assay
using online size exclusion chromatography coupled to mass spectrometry for the
assessment of protein-ligand interactions.[63] This method, named Automated Ligand
Identification System (ALIS), uses a standardized SEC-LC-MS setup, but contains an in-
house developed resin as the size exclusion material. The results obtained by ALIS have
been very impressive and next to fast affinity screening (figure 1.7, [64]) include the
recent discovery of a lipid phosphatase inhibitor involved in Diabetes type 2[65] and an
acetyl coenzyme A carboxylase ligand that is involved in antibiotics development[66].
Furthermore, it appears to be the only HTS methodology able to screen for ligands of
membrane proteins, such as G protein coupled receptors (GPCRs), which are prime
candidates to be the target of the next generation of drugs.[67]
Following the success of ALIS, a number of groups have developed innovative
applications for size exclusion affinity measurements, including some other
pharmaceutical companies. Flarakos et al. have developed a methodology able to assess
and rank ligand binding towards human serum albumin based on automated size
exclusion chromatography coupled online to a two dimensional LC-MS system, allowing
the screening of ligands with protein affinities over 3 orders of magnitude. [68] Schmidt
et al. developed a SEC-LC-MS method to analyze intact protein-arsenic complexes.[69]
Wolf et al. published a variant of SEC-LC-MS utilizing ICP-MS, especially optimized to
measure metal-ions, in this case urinary Cadmium bound to metallothionein.[70] Adam et
al. have recently expanded their earlier work in order to develop a SEC-LC-MS
methodology based on conformational changes in the protein upon binding to one of two
parnafungin isomers, thereby showing the specificity that can be accomplished using
SEC.[71]
Chapter one
31
Figure 1.7: Binding affinities calculated from ALIS data. Dissociation constants are calculated by
performing saturation experiments for 4 ligands towards muscarinic M2 acetylcholine
receptor. Affinity constants are calculated by performing displacement competition
binding experiments using the known strong binder N-methylscopolamine. Reproduced
from ref [64].
Concluding, size exclusion chromatography coupled to mass spectrometry is a very
powerful methodology, with a wide applicability, easy automation and proven results in
both fundamental research and high throughput screening. Its only disadvantages are non-
specific binding to the size-exclusion material, which can result in false positives or
negatives, and the fact that reusability of the columns is significantly reduced when more
complex protein matrices are researched.
Affinity capillary electrophoresis
Capillary electrophoresis has been used for the separation of proteins and peptides
for several decades, however, around 1992 several groups simultaneously developed
methodologies to assess protein-ligand interactions[72], or protein-DNA binding[73],
using capillary zone electrophoresis. Since that moment, the field of affinity capillary
Chapter one
32
electrophoresis (ACE) has gradually expanded and increased its applicability.[74,75]
Two major technological breakthroughs have contributed to the development of modern-
day ACE: coupling to mass spectrometry and the use of microfluidic chip technology.
Novel methodologies comprising either or both of these elements have resulted in
detection limits comparable to HPLC methodologies and proven their worth in
pharmaceutical development. When focusing on protein drug interactions, ACE
methodologies need to be separated into two categories: mobility shift methodologies and
immobilized protein methodologies.
Mobility shift methodologies
In mobility shift methodologies, the protein and ligand form a non-covalent
complex in solution. For compounds with a very slow koff, a small amount of this
incubation mixture is injected into a standard CE-MS system, and the constituents of the
mixture are separated based on the fact that the non-covalent protein ligand complex has
a distinctly different electrophoretic mobility than the non-bound protein and ligand. The
concentration of the bound and non-bound fraction can be quantified using mass
spectrometry, and protein ligand affinity can be calculated from these results. An
interesting example is published by Groesll et al. coupling CE to ICP-MS. This enabled
the researchers to assess binding between Gallium (III) based anti-cancer drugs and
serum proteins.
In the case of protein-ligand complexes with a faster koff, the complex will
dissociate quickly upon injection into the CE-MS. A small adjustment is made to
facilitate these experiments. In these cases, either the ligand or the protein will be added
to the electrophoresis buffer, thus changing the mobility of its counterpart in the
capillary. A Lineweaver-Burke plot is necessary to extrapolate the affinities from these
kinds of experiments. Many varieties of mobility shift methodologies have been
developed for various applications. The most frequently used is frontal analysis capillary
electrophoresis (FACE), distinctly different from frontal analysis chromatography in the
fact that the protein is not immobilized on a solid support, but incubated with a ligand
before injection. Recently Fermas et al. developed a methodology aimed at the study of
complexation between antithrombin and heparin pentasaccharide, a process that is
Chapter one
33
important for many biological functions.[76] They combined a continuous infusion of a
pre-incubated protein-ligand complex solution with a mass spectrometric detection. This
resulted in dramatically improved limits of detection when compared to competing MS
methodologies, allowing speculation on the future development of ligand fishing
methodologies based on FACE.
Furthermore, since the development of FACE, it has become apparent the
methodology can be used to determine protein-ligand affinity for a great variety of targets
and libraries. A small number of examples published in 2009: The interaction between
Lidocaine and hyaluronic acid[77], the interaction between a large number of drugs and
the human liposome[78] and affinity studies performed on a large testset of drugs and
human serum albumin[79]. It can safely be concluded that FACE is being routinely
employed in bioanalytical research. However, a number of improvements are still in
development, especially aimed at reducing the amount of analyte consumption. Partial
filling techniques are used to save on analyte consumption and have been around for a
little more than a decade, allowing the analysis of very expensive or complicated to
synthesize analytes.[80-82] However, these methods have a significantly higher limit of
detection than frontal analysis methodologies. A recent development to counter this
problem is partial filling multiple injection affinity capillary electrophoresis. Multiple
injections allow an electrophoretic pre-concentration in the capillary, while still using
significantly less analyte than FACE. In 2007, Zavaleta et al. developed a partial filling
multiple injection ACE methodology to study the binding of various antibiotics to D-Ala-
D-Ala-terminus peptides, thus establishing the applicability and efficiency of the
methodology.[83] More recently, Almeda et al. used a variation on this methodology to
detect antibody-antigen complexes, an application for which relatively low limits of
detection are required.[84]
Immobilized protein methodologies
Immobilized protein methodologies involve any method in which the target
protein is immobilized on a solid support, be it silica, (magnetic) beads, microfluidic
chips or any other. When a sample containing one or more possible ligands to the target
protein is injected into the system, it will bind to the immobilized target protein, thus
Chapter one
34
facilitating sample clean-up, pre-concentration, or performing a buffer change before
detection. In the past couple of years, these methodologies have especially increased in
relevance due to the relative ease of immobilizing protein on microfluidic chips and
magnetic beads. Yang et al. published an immunoaffinity CE method incorporating an
anti-�FP (�-fetoprotein) column inside a microfluidic chip.[85] This allowed them to
detect the early stage cancer-diagnostic �FP in an automated system with an analysis time
of less than one hour. Xiao et al. have published a proof-of-principle using a microfluidic
chip with immobilized strands of DNA on it, able to selectively trap insulin and insuline-
like growth factor 2, with subsequently MALDI-TOF detection.[86] This work shows
great promise for the development of a more universal microfluidic chip-based approach
for the discovery of DNA-binding ligands. The recent move towards the use of magnetic
beads in affinity measurements combined with capillary electrophoresis can also lead to
interesting results. Adachi et al. for instance, immobilized bacterial 16S rRNA on
magnetic beads, and injected them into a capillary exposed to a magnetic field, thus
trapping the beads in the capillary.[87] Then using fluorescent probes, they are able to
quantify the varying strands of rRNA, as is shown in figure 1.8. Finally, Liu et al
developed teicoplanin-derivatized microbeads, that can be immobilized inside a
microfluidic chip with microscopic magnets embedded in the PDMS chip.[88] This
allows for the calculation of the binding constant for a natively fluorescent ligand to the
antibiotic teicoplanin.
When assessing the merits of affinity capillary electrophoresis, it is apparent the
field has evolved to a point where it can compete with all other affinity methods
described in this paper. It is a relatively simple and straightforward method to implement
and use routinely. However, it is unlikely ACE will evolve into a universal technique.
Due to limitations such as non-specific adsorption of the analytes to the capillary wall,
and Joule heating of the system, many targets are not suitable for analysis by ACE,
forcing the researcher to resort to other methodologies. Furthermore, implementation of
mass spectrometric detection is not as straightforward as in liquid chromatography, and
as a result, a great variety of detection methods is used that are less specific.
Chapter one
35
Figure 1.8: The concept of immobilized magnetic bead capillary electrophoresis: (a) RNA extraction,
biotynilation and immobilization on magnetic beads. (b) injection. (c) addition of
fluorescent label. (d) charging the capillary removing all compounds not trapped
magnetically (e) injection of formamide to release the label from the target RNA. (f)
detection of the dissociated label using laser induced fluorescence.
Chapter one
36
SPR-MS
Surface Plasmon Resonance (SPR) differs fundamentally from all other methods
presented in this review, because protein binding and detection takes place at the same
location. An SPR biosensor consists of a prism positioned against a thin metal layer
(often gold) and a solution at the opposite side of the metal layer. Surface plasmons will
occur at the interface between metal and solution, and these surface plasmons have a
certain wave vector, depending on the composition and roughness of the metal surface
and the composition of the solution. When a beam of light undergoes total internal
reflection at the prism-metal interface, the photons excite the electrons in the metal film.
At one specific angle of incidence, the wave vector of these excited electrons is equal to
that of the surface plasmons, resulting in a total energy transfer. This is the definition of
surface plasmon resonance and when it happens, the light beam is no longer reflected.
The biosensor mechanism depends on the fact that the angle at which this happens is
strongly dependent on the composition of the solution. When a ligand binds to a protein
immobilized on the metal sensor, this changes the wave vector of the surface plasmons,
and the angle of incidence at which surface plasmon resonance occurs. However, no
identification occurs, as a result of which SPR is nowadays oftentimes coupled to a mass
spectrometer in order to identify the binders.
Since the launch of commercially available SPR biosensors, such as the Biacore,
in 1993[89], surface plasmon resonance has become an increasingly prominent
technology in drug discovery. In the Biacore and most competing SPR biosensors, an
analyte is infused onto a chip with immobilized target protein, starts to bind until at a
certain moment all possible binding sites are occupied, and the chip is saturated. The
speed at which this happens is dependant upon association and dissociation rates of the
complex, completely analogous to frontal affinity chromatography. The resulting data
and data processing is also analogous to FAC. However, in SPR, the change in signal is
not dependant upon analyte concentration, but analyte molecular weight. This is why
SPR bioassays are especially suitable for the study of protein-protein interactions, or
other large molecules that bind to the immobilized target protein.
Chapter one
37
The first major applications of SPR biosensors were in the field of antibody-
antigen interactions[90], the study of DNA hybridization and dehybridization,[91] and
many other macromolecular interactions.[92] Around the same time, the first coupling of
SPR to MALDI-TOF mass spectrometry was reported, in which the SPR chip was
directly used as MALDI chip.[93,94]. Since that moment, many SPR-MALDI
methodologies have been developed, covering a wide assortment of target proteins.[95-
98]. A number of interesting reviews have been published providing an overview of the
importance of SPR-MS for modern-day proteomics.[99,100]
Interesting papers in recent SPR-MS method development include the work of
Borch et al. who exposed an immobilized enzyme to a possible inhibitor, and proceeded
to perform an activity assay by infusing the substrate and analyzing the substrate
conversion by MS.[101] Bouffartigues et al. published a high-throughput SPR-SELDI-
MS methodology able to identify protein bound to DNA.[102] Marchesini et al.
published an SPR methodology coupled online to a nano-HILIC-LC system, resulting in
an online electrospray interface between the SPR-LC and the mass spectrometer.[103]
In the field of small molecule research, significantly fewer methodologies have
been published, presumably due to the dependency of the SPR signal on molecular
weight and as a result, its improved performance for analytes above 15 kDa. An elegant
solution for analyzing small molecules is to immobilize the small molecules on the chip,
and subsequently measuring the binding of the target protein,[104] analogous to
immobilized antigens.[90] However, recently Jecklin et al. developed an SPR
methodology for seven analytes under 500 Da. Utilizing human carbonanhydrase
immobilized on the SPR chip, they managed to calculate accurate dissociation constants
for the testset, and compare them to two reference methods.[103] Also, Marchesini et al
successfully measured the binding and dissociation kinetics of a testset of seven low
molecular weight paralytic shellfish poisons binding to immobilized saxitoxin
monoclonal antibody (Figure 1.9).[105] Furthermore, recent publications by Wang et al.
and Miura et al. show the development of indirect SPR affinity methods, in which the
affinity of a small molecule is determined by its competition with a large ligand already
bound to the immobilized target protein.[106,107]
Chapter one
38
Figure 9: Binding and dissociation kinetics of seven paralytic shellfish poisons to
immobilized MAb GT13A. Reproduced from ref [105].
Concluding, SPR is an extremely powerful tool for protein affinity analysis, mainly
because of the quantification of the affinity of the analyte during it’s immobilization on
the SPR chip-surface. Its main drawback is that it is most efficient for compounds with a
high molecular weight, and thus most suitable for the study of protein-protein
interactions. However, developments that target small molecules are upcoming.
Conclusion
Reviewing the wide spectrum of methodologies discussed in this paper, all of
them can be separated into two general categories, with distinctly different applicability,
advantages and disadvantages. The first group consists of methods that immobilize the
target protein on a solid support, the second of methodologies that allow in solution
complexation. Ultrafiltration and size exclusion are solution based methods. The complex
is formed in solution, approaching in vivo circumstances as closely as possible. The same
goes for dynamic protein affinity selection and a number of capillary electrophoresis
Chapter one
39
methods. All other techniques: frontal affinity, zonal elution, affinity capture, most CE-
based methods and surface plasmon resonance use immobilized proteins.
When assessing the progress made using immobilized proteins, a number of
conclusions can be drawn. Frontal affinity chromatography will remain relevant, because
it is a cheap and easy to implement method to assess protein-ligand binding. The
immobilized target proteins are getting more and more complex, and none of the in-
solution based methods rivals the data resulting from FAC methodologies. The only
methodology that does is SPR. SPR-MS can produce the same affinity data as FAC, and
probably more. However, SPR is limited by its inability to measure small ligands
directly, and the more limited amount of target proteins that have proven to be
successfully immobilized on the SPR chip.
The solution-based methodologies might not provide the same amount of affinity
data, but they do not suffer from impaired binding properties due to protein
immobilization, and each of them has advantages of its own over the competition.
Ultrafiltration benefits from the reusability of the receptor, and the speed of
measurement. Capillary electrophoresis affinity selection can target proteins unsuitable
for any of the other methodologies, and will stay important despite issues with non-
specific absorption of the proteins to the capillary wall. Dynamic protein affinity
selection is very sensitive for weak binders, and suffers little from non-specific binding.
However, the size exclusion methodologies are the ones most commonly applied in
modern drug discovery for two simple reasons. SEC is very easy to implement and
automate using standard 96 well plates and equipment, and SEC is significantly faster
than all other techniques discussed.
As a result, when reviewing literature or visiting relevant conferences, most of the
research on affinity selection mass spectrometry in pharmaceutical industry is focused on
surface plasmon resonance technologies and size exclusion technologies. However, many
if not all of the discussed fields of affinity selection mass spectrometry will undoubtedly
result in new lead candidates in the upcoming years.
Chapter one
40
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