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Supplement tooctober 2015
www.chromatographyonline.com
Biopharmaceutical Analysis
Advances in
ES683229_LCESUPP1015_CV1.pgs 10.01.2015 20:41 ADV blackyellowmagentacyan
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ES683633_LCESUPP1015_CV2_FP.pgs 10.01.2015 22:43 ADV blackyellowmagentacyan
3www.chromatographyonline.com
6 Introduction Pat Sandra and Koen Sandra
An introduction from the guest editors of this special supplement.
8 Modern Column Technologies for the Analytical Characterization of Biopharmaceuticals in Various Liquid Chromatographic Modes
Szabolcs Fekete, Jean-Luc Veuthey, and Davy GuillarmeThe recent trends in column technology for reversed-phase LC, SEC, IEX, and HIC for analysis of biopharmaceuticals at the protein level is critically discussed.
16 Monoclonal Antibodies and Biosimilars — A Selection of Analytical Tools for Characterization and Comparability Assessment Koen Sandra, Isabel Vandenheede, Emmie Dumont, and Pat SandraWith the top-selling mAbs evolving out of patent there has been a growing interest in the development of biosimilars. In demonstrating comparability to the originator product, biosimilar developers are confronted with an enormous analytical challenge. This article presents a selection of state-of-the-art analytical tools for mAb characterisation and comparability assessment.
24 Harnessing the Benefi ts of Mass Spectrometry for In-depth Antibody Drug Conjugates Analytical Characterization
Alain Beck and Sarah CianféraniRecent progress in high-resolution mass spectrometry (HRMS) and liquid chromatography–mass spectrometry (LC–MS) methods for the structural characterization of brentuximab vedotin and trastuzumab emtansine are presented.
31 Higher Order Mass Spectrometry Techniques Applied to Biopharmaceuticals
Christian G. HuberAn outline of the basic principles of MS techniques used to investigate higher order structural features of biopharmaceuticals, as well as some insights into applications relevant to the pharmaceutical industry.
38 Advances in Liquid Chromatography–Tandem Mass Spectrometry (LC–MS–MS)-Based Quantitation of Biopharmaceuticals in Biological Samples
Nico C. van de MerbelThe technical requirements for a successful LC–MS–MS method for the quantitation of biopharmaceuticals are presented and the advantages and disadvantages compared to ligand-binding assays are evaluated.
45 Analyzing Host Cell Proteins Using Off-Line Two-Dimensional Liquid Chromatography–Mass Spectrometry
Koen Sandra, Alexia Ortiz, and Pat SandraThe use of off-line 2D-LC–MS for the characterization of HCPs and their monitoring during downstream processing.
Advances in
Biopharmaceutical Analysis
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4 Advances in Biopharmaceutical Analysis – October 2015
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Pat SandraResearch Institute for Chromatography,
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The Netherlands
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ES683551_LCESUPP1015_005_FP.pgs 10.01.2015 21:54 ADV blackyellowmagentacyan
When we were asked to edit a follow-up to
the LCGC Europe May 2013 supplement
“Advances in Pharmaceutical Analysis”,
we immediately wanted to highlight the
challenges in biopharmaceutical analysis.
Indeed, within the pharmaceutical industry
and also within our own research activities
related to pharmaceutical analysis, there
has been a remarkable shift from small to
large molecules. On the market since the
early 1980s, protein biopharmaceuticals
have seen an enormous growth in the last
decade. It is even expected that within
the current decade, more than 50% of
new drug approvals will be biological in
nature. A dominant role is thereby played
by monoclonal antibodies (mAbs) of
which a substantial number have reached
blockbuster status. The top-ten bestselling
pharmaceuticals are currently heavily
populated by mAbs.
Protein biopharmaceuticals are
large and heterogeneous and their
in-depth analysis during development
and also during their lifetime requires
the best of both chromatography and
mass spectrometry (MS). In editing this
special issue, we have therefore selected
authorities in the field to illustrate the
state-of-the-art in biopharmaceutical
analysis.
The first contribution, authored by
Szabolcs Fekete, Jean-Luc Veuthey, and
Davy Guillarme, provides an overview
of the different LC column formats
recently introduced in the market for
reversed-phase, size-exclusion (SEC),
ion-exchange (IEX), and hydrophobic
interaction (HIC) chromatographic
analyses of therapeutic proteins, mAbs,
and antibody-drug-conjugates (ADCs).
In the May 2013 supplement we
described the features of liquid
chromatography coupled to mass
spectrometry (LC–MS) in the
characterization of protein
biopharmaceuticals. With the
patents of the first generation protein
biopharmaceuticals expired and
blockbuster mAbs appearing on the
market, activities in biosimilars have
exploded in recent years. More than
15 biosimilars have already been
approved in Europe while a version
of filgrastim will be launched in the
U.S. as the first biosimilar towards the
end of 2015. Analytical methods to
compare originators with biosimilars are
highlighted in the second contribution
from our team at the Research Institute for
Chromatography.
The antibody market has been
reshaped by various next-generation
formats (bio specific mAbs, antibody
mixtures, nanobodies, brain penetrant
mAbs, glyco-engineered formats), and
in recent years the ADCs brentuximab
vedotin and trastuzumab emtansine
have been approved by the EMA and
the FDA. In ADCs a cytotoxin is coupled
to an antibody that specifically targets a
certain tumour marker. As such, highly
toxic drugs can be delivered in a targeted
fashion to tumour cells without affecting
healthy cells. Compared to naked mAbs,
the conjugation of cytotoxic drugs further
adds to the complexity. The power of MS
to unravel this complexity is illustrated in
the second paper authored by Alain Beck
and by Sarah Cianferani.
The previous two contributions clearly
illustrate the importance of MS in the
elucidation of the primary structure
of therapeutic proteins. Higher order
elements, on the other hand, can be
derived from special MS technologies
such as native MS, ion mobility MS,
hydrogen-deuterium exchange MS,
and chemical cross-linking MS. In the
fourth contribution, Christian Huber
describes the basic principles of
these techniques and illustrates their
features for the characterization of
higher order structures of some protein
biopharmaceuticals.
Traditionally, ligand-binding assays
(LBAs) are applied to study the
pharmacokinetic behaviour of protein
biopharmaceuticals in biological
fluids. LBAs are characterized by a
high throughput and sensitivity but
may suffer from long development
times and potential interferences from
other proteins present in the matrix. In
addition, generation of drug specific
antibody tools is a time-consuming
process. Liquid chromatography
coupled to tandem mass spectrometry
(LC–MS–MS) methods are more and
more used as alternatives to LBAs,
often offering improved figures-of-merit
while at the same time being generically
applicable. Some of the technicalities and
advantages and disadvantages of LC–
MS–MS compared to LBAs for monitoring
biopharmaceuticals in biological fluids are
addressed in the fifth contribution by Nico
C. van de Merbel.
The presence of residual host cell
proteins (HCPs) is a potential safety
risk in any biopharmaceutical product.
Despite enormous purification efforts,
these HCPs may be left behind from the
expression hosts. HCPs are normally
dosed during downstream processing
and in the final biopharmaceutical product
by enzyme-linked immunosorbent
assays (ELISA). As mentioned in the
previous paper, LBAs are more and more
complemented or even replaced by LC–
MS–MS and this is illustrated in the last
contribution by our group. The use of off-
line two-dimensional LC–MS–MS in the
characterization of HCPs is described and
the added value of using multidimensional
chromatography is clearly demonstrated.
We hope that the contributions in this
supplement are of interest and even a
source of inspiration to the numerous
analysts in the (bio)pharmaceutical
industry. It was a pleasure for us to
edit and review the contributions of
outstanding (preselected) colleagues.
We would like to thank all of them for
their excellent work.
Advances in Biopharmaceutical AnalysisPat Sandra and Koen Sandra, Research Institute for Chromatography, Kortrijk, Belgium.
An introduction from the guest editors of this special supplement from LCGC Europe focusing on recent developments in biopharmaceutical analysis.
6 Advances in Biopharmaceutical Analysis – October 2015
Pat Sandra Koen Sandra
ES683465_LCESUPP1015_006.pgs 10.01.2015 21:10 ADV blackyellowmagentacyan
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ES683547_LCESUPP1015_007_FP.pgs 10.01.2015 21:53 ADV blackyellowmagentacyan
Therapeutic proteins are large
and heterogeneous molecules
subjected to a variety of enzymatic
and chemical modifications
during expression, purification,
and long‑term storage. These
changes include several possible
modifications, such as oxidation,
deamidation, glycosylation,
aggregation, misfolding, or
adsorption, leading to a potential loss
of therapeutic efficacy or unwanted
immune reactions. Regulatory bodies
require a detailed characterization
(for example, verifying primary
structure and appropriate
post‑translational modifications,
secondary and tertiary structure),
lot‑to‑lot and batch‑to‑batch
comparisons, stability studies,
impurity profiling, glycoprofiling,
determination of related proteins and
excipients as well as determination of
protein aggregates. For this purpose,
a single analytical technique is
generally not sufficient, and a variety
of orthogonal methods are required
to fully describe such a complex
sample.
Today, one of the most widely
used analytical techniques for
therapeutic protein characterization
is liquid chromatography (LC). This
is probably a direct result of the
remarkable developments of the
past few years, which have enabled
a new level of chromatographic
performance. These developments
include ultrahigh‑pressure LC
(UHPLC), columns packed with
wide‑pore superficially porous
particles (SPPs), and organic
monolith columns, which have
allowed a dramatic increase in
separation efficiency, even with large
intact biomolecules.
This article will review the
possibilities and trends of current
state‑of‑the‑art LC column
technology applied for different
modes of chromatography for the
characterization of therapeutic
proteins.
Hydrophobic Interaction ChromatographyHydrophobic interaction
chromatography (HIC) has been
historically used for protein
purification; more recently, the
two main application fields have
been in the determination of the
drug‑to‑antibody ratio (DAR) of
antibody‑drug conjugates (ADCs)
and in monitoring post‑translational
modifications of monoclonal
antibodies (mAbs).
In HIC, proteins are retained and
separated on the basis of their
hydrophobicity as a result of the
van der Waals forces between the
hydrophobic ligands of the stationary
phase and the non‑polar regions of
proteins (1). The binding of proteins
to a hydrophobic surface is affected
by a number of factors including the
type of ligand, the ligand density
on the solid support, the backbone
material of the stationary phase, the
hydrophobic nature of the protein,
and the type of salt added to the
mobile phase. During the separation,
a negative salt gradient (typically
from 2–3 M to 0 M) is applied under
aqueous mobile phase at around
pH 6.8–7.0. The structural damage
to the biomolecules is therefore
minimal and its biological activity is
maintained (2).
Analytical‑scale HIC columns
are based either on silica or
polymer particles. Both porous and
non‑porous particles are available.
Highly cross‑linked non‑porous
poly(styrene–divinylbenzene) (PS/
DVB) and polymethacrylate‑based
particles are frequently used in
protein separations as a result of
their advantageous mass transfer
properties (the main contribution
to the band broadening of large
biomolecules, namely trans‑particle
mass transfer resistance is
negligible). Table 1 summarizes the
most widely used and the latest HIC
columns applied for mAb and ADC
separations.
These materials can now withstand
pressure drops of up to 100–400
bar. Columns are typically packed
with 10‑, 7‑, 5‑, 3‑, and even 2.5‑µm
particles. Column diameters between
2 mm and 8 mm are available
but 4.6‑mm i.d. columns are the
most widely used in current HIC
applications. It is worth mentioning
that there is a need for 150 × 2.1
mm column formats, which are often
applied for the analysis of proteins in
modern chromatographic practice.
Modern Column Technologies for the Analytical Characterization of Biopharmaceuticals in Various Liquid Chromatographic Modes Szabolcs Fekete, Jean-Luc Veuthey, and Davy Guillarme, School of Pharmaceutical Sciences, University of Geneva,
University of Lausanne, Geneva, Switzerland.
The recent trends in column technology for reversed-phase liquid chromatography (LC), size-exclusion chromatography (SEC), ion-exchange chromatography (IEX), and hydrophobic interaction chromatography (HIC) for analysis of biopharmaceuticals at the protein level is critically discussed.
Ph
oto
Cre
dit: Jo
rg G
reu
el/G
ett
y Im
ag
es
8 Advances in Biopharmaceutical Analysis – October 2015
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9www.chromatographyonline.com
Guillarme et al.
Table 1: Recent state‑of‑the‑art and some widely used “reference” columns applied for the separation of therapeutic proteins in HIC, SEC,
IEX, and reversed‑phase LC modes.
Column Name ChemistryParticle size/Macropore Size (µm)
Max Temperature
(oC)pH Range
Max Pressure
(bar)
HIC Columns
TSKgel (Tosoh)
Butyl‑NPR C4 (non porous) 2.5
50 2–12
200
Ether‑5PW Ether (porous) 1050
Phenyl‑5PW Phenyl (porous) 10
Protein‑Pak Hi Res HIC (Waters) C4 2.5 60 2–12 200
Thermo
MAbPac HIC‑Butyl C4 5 60 2–12 300
MAbPac HIC‑20 Alkylamide 5 60 2–9 400
ProPac HIC‑10 Amide/ethyl 5 60 2.5–7.5 300
IEX Columns
Proswift (Thermo)
(monolith)
SAX‑1SStrong anion exchange
(quaternary amine)
Information
not
available
70
2–12 70
WAX‑1SWeak anion exchange
(tertiary amine)60
WCX‑1SWeak cation exchange
(carboxylic acid)60
SCX‑1SStrong cation exchange
(sulphonic acid)60
TSKgel (Tosoh)
SCXStrong cation exchange
(sulphonic acid)5
45
2–14
50
SuperQ‑5PWStrong cation exchange
(trimethylamino)10 2–12
SP‑STATStrong cation exchange
(sulphopropyl)7, 10 3–10
Q‑STATStrong anion exchange
(quaternary ammonium)7, 10 3–10
Bio Mab (Agilent)Weak cation exchange
(carboxylate)
1.7 3 5 10
80 2–12
270 410 550 680
Antibodix (Supelco, Sepax)Weak cation exchange
(carboxylate)
1.7 3 5 10
80 2–12
270 410 550 680
Protein‑Pak Hi Res
IEX (Waters)
SPStrong cation exchange
(sulphopropyl)7
60 3–10
100
CM Weak cation exchange
(carboxymethyl)7 100
QStrong anion exchange
(quaternary ammonium)5 150
MAbPac SCX‑10 (Thermo)Strong cation exchange
(sulphonic acid)
3 5 10
60 2–12480 480 200
Bio‑Pro (YMC)
QA
QA‑F
Strong anion exchange
(quaternary ammonium) 5 60 2–12
30 120
SP
SP‑F
Strong cation exchange
(sulphopropyl)30 120
SEC Columns
Thermo Silica‑based 3 60 2.5–7.5 200
YMC‑Pack Diol‑SEC Diol modified silica‑based 5 40 5–7.5 200
Acclaim SEC‑300 (Thermo) Hydrophilic polymethacrylate resin 5 60 2–12 1200
TSKgel SW aggregate (Tosoh) Diol 3 30 2.5–7.5 120
TSKgel SW mAb (Tosoh) Diol 4 30 2.5–7.5 120
SRT‑SEC (Sepax) Surface‑coated silica‑based 5Information not
available2–8.5
Information
not available
Zenix‑SEC (Sepax) Surface‑coated silica‑based 3 ~250 2–8.5 80
ES683258_LCESUPP1015_009.pgs 10.01.2015 20:46 ADV blackyellowmagentacyan
HIC allows both the characterization
of the distribution of drug‑linked
species and the determination of
average DAR of ADCs (3). Conjugation
of the drug‑linker to the antibody
increases the hydrophobicity;
therefore HIC appears as a suitable
tool to separate the different DAR
species. A good example of the HIC
profile of a native IgG1 ADC is shown
in Figure 1 (4).
Recently an off‑line mass
spectrometric (MS) detection was
applied for the characterization of
Brentuximab‑vedotin. Each individual
HIC peak was collected, buffer
exchanged, and analyzed by native MS
(5). HIC was also successfully applied
for monitoring various post‑translational
modifications, including proteolytic
fragments, domain misfolding,
tryptophan oxidation, and aspartic acid
isomerization in therapeutic mAbs (6).
Ion-Exchange ChromatographyIon‑exchange chromatography (IEX)
is widely used for the characterization
of therapeutic proteins and can
be considered as a reference
marker and powerful technique
for the qualitative and quantitative
evaluation of charge heterogeneity.
Among the different IEX modes,
cation‑exchange chromatography
(CEX) is the most widely used for
protein characterization (7).
Two modes of elution are often
applied for protein characterization,
namely the (i) salt‑gradient and the
(ii) pH‑gradient. In salt‑gradient
mode, solutes are eluted in order of
increasing binding charge, which
correlates more or less with the
isoelectric point (pI) and equilibrium
constant. In this case, the mobile
phase pH is kept constant, while
the ionic strength is continuously
increased. In pH‑gradient mode,
the ionic strength is kept constant
and the pH is varied during the
gradient programme. This mode
of elution is often referred to as
chromatofocusing.
10 Advances in Biopharmaceutical Analysis – October 2015
Guillarme et al.
Table 1: Contd....
Column Name ChemistryParticle Size/Macropore Size (µm)
Max Temperature
(oC)pH Range
Max Pressure
(bar)
Bio SEC (Agilent) surface‑coated silica‑based3 Information not
available2–8.5 240
5
Acquity UPLC BEH SEC (Waters) diol modified hybrid‑based1.7
60 2–8 6002.5
Reversed-phase LC Columns
ProSwift (Thermo)
(Monolith)
RP‑1S
Phenyl
1 70 1–14 200
RP‑2H 2.2 70 1–14 200
RP‑3U 5.1 70 1–14 200
RP‑10RInformation
not available80 1–10 300
Acquity BEH 300 (Waters) C18, C4 1.7 80 1–12 1000
Zorbax (Agilent)
300SB RRHD C18, C8 1.8 80 1–8 1200
Poroshell SB300 C18, C8, C35 (0.25‑µm
thickness)90 1–8 600
Poroshell 300Extend C185 (0.25‑µm
thickness)60 2–11 600
AdvanceBio
RP‑mAbC8, C4, diphenyl
3.5 (0.25‑µm
thickness)90 1–8 600
Aeris
(Phenomenex)
Widepore C18, C8, C43.6 (0.2‑µm
thickness)
90 (C18,C8),
60 (C4)1.5–9 600
Peptide C18
3.6 (0.5‑µm
thickness)
2.6 (0.35 µm
thickness)
1.7 (0.22‑µm
thickness)
90 1.5–9
600
1000
Halo (Advanced
Materials
Technology)
Peptide C18, CN
2.7 (0.5‑µm
thickness)
4.6 (0.6 µm
thickness)
100 1–9 600
Protein C18, C83.4 (0.2‑µm
thickness)90 1–9 600
Flare Widepore (Diamond Analytics) C183.6 (0.1‑µm
thickness)100 1–13 400
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Regarding the stationary phase,
there are two main aspects that need
to be considered for successful
IEX separation: (i) the strength
of interaction and associated
retention (strong or weak ion
exchanger) and (ii) the achievable
peak widths (efficiency) (8). Both
cation and anion exchangers can
be classified as either weak or
strong exchangers. Weak cation
exchangers are comprised of a weak
acid that gradually loses its charge
as the pH decreases (for example,
carboxymethyl groups), while strong
cation exchangers are comprised of
a strong acid that is able to sustain
its charge over a wide pH range
(for example, sulphopropyl groups).
On the other hand, strong anion
exchangers contain quaternary
amine functional groups, while
weak anion exchanger possesses
diethylaminoethane (DEAE) groups.
As a rule of thumb, it is preferable to
begin the method development with
a strong exchanger, to ensure that a
broad pH range can be worked on.
Strong exchangers are also useful if
the maximum resolution occurs at an
extreme pH. However, silica‑based
ion exchangers can only be operated
in a restricted pH range. In contrast,
polymeric ion exchangers can be
used over a wide pH range.
Commercially available
IEX columns are based on
silica or polymer particles but
organic‑polymeric monoliths are
also available. Both porous and
non‑porous particles are available
but for large molecules, which
possess low diffusivity, non‑porous
materials are clearly preferred. Highly
cross‑linked non‑porous PS/DVB
materials are most frequently used in
protein separations because of their
high pH stability (2 ≤ pH ≤ 12). These
materials can now withstand pressure
drop of up to 500–600 bar in some
cases. Columns packed with 10‑,
5‑, or 3‑µm non‑porous particles are
often used, but sub‑2‑µm materials
are also available to perform UHPLC
separations (see Table 1). Suitable
peak capacity can be attained with
large biomolecules on those columns
within a reasonable analysis time (for
example, 15–20 min). However, some
limitations can be expected in terms
of loading capacity and retention
when applying non‑porous materials.
A recent study systematically
compared the latest state‑of‑the‑art
cation exchanger columns applied
for the characterization of therapeutic
mAbs in pH‑ and salt‑gradient modes
(8).
Figure 2 shows an example of the
separation of four intact antibody
charge variants using a 100 × 4.6
mm, 5‑µm strong cation polymeric
exchanger column packed with
non‑porous particles and a 20‑min
long gradient (9)
Size-Exclusion ChromatographySize‑exclusion chromatography
(SEC) is a powerful technique for
the qualitative and quantitative
evaluation of protein aggregates. The
main advantage of SEC is the mild
mobile phase conditions that permit
the characterization of proteins with
minimal impact on the conformational
structure and local environment.
SEC separates biomolecules
according to their hydrodynamic
radius. The stationary phase consists
of spherical porous particles with a
carefully controlled pore size, through
which the biomolecules diffuse based
on their molecular size difference
using an aqueous buffer as the
mobile phase. Basically, SEC is an
entropically controlled separation
process in which molecules are
separated on the basis of molecular
size differences (filtering) rather
than by their chemical properties
(10). Therefore, retention factor
12 Advances in Biopharmaceutical Analysis – October 2015
Guillarme et al.
200
150
100
50
1210
DAR 4
DAR 6
Time (min)
Sig
na
l
DAR 8
DAR 2
DAR 0
864
0
Figure 1: HIC separation of an ADC for the determination of drug‑to‑antibody ratio (DAR). Adapted and reproduced with permission from Analytical Chemistry 84, Lan N. Le, Jamie M.R. Moore, Jun Ouyang, et al., Profiling Antibody Drug Conjugate Positional Isomers: A System-of-Equations Approach, 7479–7486 (2012) © American Chemical Society.
Sig
nal
2015
Time (min)
1050
1 2 3 4
Figure 2: IEX separation of four intact mAbs (natalizumab [1], cetuximab [2], adalimumab [3], and denosumab [4]). Adapted and reproduced with permission from Journal of Pharmaceutical and Biomedical Analysis 111, Szabolcs Fekete, Alain Beck, and Davy Guillarme, Characterization of cation exchanger stationary phases applied for the separations of therapeutic monoclonal antibodies, 169–176 (2015) © Elsevier.
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(thermodynamic) in SEC is different from other chromatographic modes. Here, the thermodynamic retention factor is the fraction of the intraparticle pore volume that is accessible to the analyte (11).
Since no retention occurs in SEC, large pore volumes (high porosity) are required to ensure appropriate resolution. Generally, this large pore volume is provided by long‑and wide‑bore columns. In routine SEC applications, a 30‑cm column length with internal diameters (i.d.) of 7.8‑, 8.0‑, or 10‑mm is generally employed. These SEC columns are referred to as standard bore columns. Now, several vendors offer narrow bore columns with 4.6‑mm i.d. and 15‑cm length that are packed with very efficient, small particles of ~3 µm. Similar separation power can be attained using these columns as with 5‑µm particles in 30‑cm standard bore columns, but the analysis time can be reduced by a factor of 3 to 4 (12).
There are mainly two types of SEC packing materials: (i) silica, with or without surface modification, and (ii) cross‑linked polymeric packings, which possess non‑polar (hydrophobic), hydrophilic, or ionic character (10). The most common silica packing consists of chemically bonded 1,2‑propanediol functional groups that provide a hydrophilic surface. This stationary phase blocks or reacts with many of the acidic silanol groups allowing the surface to be neutralized. Bare silica is also a suitable packing material for non‑aqueous polar or non‑polar organic mobile phases; however, it is not recommended with aqueous mobile phases because of the presence of active silanol sites. The latest type of silica‑related packing is an ethylene‑bridged hybrid inorganic‑organic (BEH) material that is currently available at particle sizes of 1.7 µm — the first sub‑2‑µm SEC packing — and 2.5 µm (13). Compared to regular silica packings, BEH particles have improved chemical stability as well as reduced silanol activity. The 1.7‑µm BEH material can be operated at up to 600 bar.
There have been a number of different hydrophilic cross‑linked packings developed for the SEC of
biopolymers. Most of these packings are proprietary hydroxylated derivatives of cross‑linked polymethacrylates (10). Unusual polymeric packings for aqueous SEC include sulphonated cross‑linked polystyrene, polydivinylbenzene derivatized with glucose or anion exchange groups, a polyamide polymer, and high‑performance, crossed‑linked agarose (10).
Today, columns for aqueous and non‑aqueous SEC applications with pore sizes of 125 to 900 Å are commercially available (14). Very fast separations of peptides, myoglobin, and insulin aggregates have been demonstrated with 1.7‑µm SEC columns (15). These columns were also applied for the characterization of recombinant mAbs (13).
Applying 1.7‑ and 2.5‑µm particles in SEC has opened up a new level of separation performance, but it should be kept in mind that on very fine particles, the separation quality is improved at the cost of pressure (and frictional heating temperature gradients). Therefore, there is a risk of creating on‑column aggregates when analyzing sensitive proteins under high pressure (> 200 bar) conditions (13).
Reversed-Phase Liquid Chromatography In reversed‑phase liquid chromatography (LC), the solute retention is predominantly mediated through hydrophobic interactions between the non‑polar amino
acid residues of the proteins and the bonded n‑alkyl ligands of the stationary phase. Compared to the HIC mode, the reversed‑phase LC mobile phase typically consists of water, acetonitrile or methanol, and 0.1–0.2% trifluoroacetic acid or formic acid. The separation mechanism is based on a combination of solvophobic and electrostatic interactions, the latter being governed by the interaction of TFA with basic side chains of a few amino acids (that is, arginine, lysine, and histidine) and the N‑terminus as well as ionic interactions between the positive charges at the surface of the protein and the negatively charged residual silanols (16). The efficiency of reversed‑phase LC is always superior to other chromatographic modes and its superior robustness makes it well suited for use in a routine environment (17).
Current reversed‑phase LC stationary phases used for proteins analysis can be classified as silica‑based particulate materials and organic monoliths. The pore size of particulate phases is an important factor that must be considered. For the analysis of peptides and small proteins, a pore size between 100–200 Å may be acceptable. However, porous materials with pore sizes of more than 200 Å are mandatory for the separation of larger proteins or mAbs fragments because the solute molecular diameter must be approximately one‑tenth the size
13www.chromatographyonline.com
Guillarme et al.
64 5
Time (min)
31 20
Sig
nal
L H
HC
2xLC2xHC
Reduction
+
Figure 3: Reversed phase LC analysis of reduced IgG1 mAb. Unpublished results from the authors’ laboratory.
ES683253_LCESUPP1015_013.pgs 10.01.2015 20:46 ADV blackyellowmagentacyan
of the pore diameter to avoid the restricted diffusion of the solute and to allow the total surface area of the sorbent material to be accessible. An average pore size between 250–300 Å is often mentioned as the reference value for protein separations, but recently it was shown that 400 Å particles completely eliminated restricted diffusion effects for molecules up to about 500 kDa.
The two main trends today in reversed‑phase LC analysis of therapeutic proteins are the use of (i) fully porous small particles (FPPs) (sub‑2‑µm) and (ii) superficially porous particles (SPPs), which possess particle sizes between 3‑ and 4‑µm.
Columns packed with FPPs have constraints in separation speed and efficiency because of limitations in the stationary phase mass transfer, which results from the relatively long diffusion times required for proteins to cross the porous structure. Therefore, Horváth first applied the concept of SPPs in the late 1960s (18,19). They were initially intended for the analysis of macromolecules such as peptides and proteins. SPPs are made of a solid, non‑porous silica core surrounded by a porous shell layer. They have similar properties to the fully porous materials conventionally used in HPLC. The rationale behind this concept was to improve column efficiency by shortening the diffusion path that molecules must travel, in addition to improving their mass transfer kinetics.
It was recently shown that columns
packed with wide‑pore 3.6‑µm and 3.4‑µm SPPs showed significant gain in analysis time and peak capacity compared to FPPs for intact protein analysis (20,21). These wide‑pore SPPs are now available with C4, C8, and C18 chemistries and can be operated up to 600 bar. Figure 3 shows an example of fast separation of heavy‑chain (Hc) and light‑chain (Lc) variants of an IgG1 mAb performed on a wide‑pore C4 SPP column.
In another study, efficiency and analysis times of 1.7‑µm SPPs and FPPs were compared for peptides and moderate size intact proteins (22). This study suggests a two‑fold increase in terms of achievable peak capacity and analysis time for large proteins when using SPPs compared to FPPs of the same size. For the separation of peptides and moderate size proteins, a 160 Å SP packing was also introduced (23,24). Recently 1.3‑ and 1.6‑µm SPPs were also applied for peptide mapping of mAb samples (25,26). By combining long columns (200–300 mm) with extended analysis time, peak capacity around 1000 can be reached with 1.3‑µm SPPs for 0.5–2 kDa peptides.
An alternative, carbon‑nano‑diamond based C18 superficially porous material was recently introduced (27). The core of this material is a carbonized poly(divinylbenzene) particle with a diameter of approximately 3.4‑μm. Poly(allylamine)‑nanodiamond hetero‑layers are deposited onto the surface of the carbonized core by a modified layer‑by‑layer method. The resulting core‑shell is synthesized to
a shell thickness of ca. 0.1 μm and a finished particle size of 3.6 μm. This superficially porous carbon‑based material was successfully applied for real life protein separations.
Another interesting alternative to SPPs was proposed by Hayes et al. (28). The so‑called sphere‑on‑sphere (SOS) approach provides a simple and fast one‑pot synthesis in which the thickness, porosity, and chemical substituents of the shell can be controlled by using the appropriate reagents and conditions (29). SOS particles have been shown to be microporous with a pore diameter of less than 2 nm. However, while the surface of the material might not exhibit significant porosity, when packed into a HPLC column the spaces between surface nanospheres provide superficial macroporosity. It has been proposed that for large molecules, larger pores as well as reduction of the shell thickness can be advantageous, because of the shorter diffusion distance and greater access to the surface area of the material (30). SOS particles were demonstrated to have similar chromatographic performance compared to commercial SPP materials (28). Figure 4 shows the separation of reduced ADC (Brentuximab‑Vedotin) fragments on a column packed with SOS particles.
As an alternative to particle‑based stationary phase formats for the LC separations of proteins, organic polymer‑based monoliths offer some advantages, including high permeability and rapid mass transfer (31). Polymeric monolithic stationary phases have shown great potential for the reversed‑phase LC separations of large biomolecules, including intact proteins, oligonucleotides, and peptides. With this material, the mass transfer is mainly driven by convection, rather than diffusion, because of the absence of mesopores (32). The fact that the solvent is forced to pass through the macropores of the polymer because of pressure leads to faster convective mass transfer compared to the slow diffusion process into the stagnant pore liquid that is present in porous beads packed columns. As a result of their open channel structure, monoliths generally possess a high
14 Advances in Biopharmaceutical Analysis – October 2015
Guillarme et al.
Sig
na
l
128 10
Time (min)62 40
H1
H0
L1L0
H2H3
Figure 4: Reversed phase LC separation of reduced ADC (Brentuximab‑Vedotin) fragments. Unpublished results from the authors’ laboratory.
ES683259_LCESUPP1015_014.pgs 10.01.2015 20:46 ADV blackyellowmagentacyan
permeability, allowing the application of elevated flow rates at moderate back pressure. It was previously demonstrated that polymeric stationary phases led to superior performance over silica‑based materials in the reversed phase analysis of very large proteins (MW >50 kDa) (33).
ConclusionThere is always a need to use several chromatographic methods to draw reliable conclusions regarding the quality of biopharmaceuticals. IEX, SEC, and HIC are historical techniques and are still used in any laboratory dealing with the analytical characterization of mAbs or ADCs. These techniques were known to offer poor resolving power, which is why the stationary phases employed in IEX, SEC, and HIC have strongly evolved over the last few years, in terms of chemistries, dimensions, and chemical stability.
The most important improvements for protein analysis were brought to reversed‑phase LC materials. In the past, this technique has rarely been used for biopharmaceutical characterization. However, because this is the only chromatographic approach directly compatible with MS, providers have improved and developed their existing materials. The performance that can be achieved today with columns packed with wide pore sub‑2‑µm fully porous or sub‑4‑µm superficially porous are highly competitive, and even if the selectivity of reversed‑phase LC is still limited for separating charge or size variants of proteins, this is (at least partially) compensated by the high kinetic performance generated by modern reversed‑phase LC columns.
AcknowledgementsThe authors acknowledge Alain Beck (Pierre Fabre, Saint‑Julien Genevois, France) for providing mAb and ADC samples, and Stephanie Schuster (Advanced Materials Technology) and Tony Edge and Richard Hayes (Thermo Fisher Scientific) for providing stationary phases.
Davy Guillarme wishes to thank the Swiss National Science Foundation for support through a fellowship to Szabolcs Fekete (31003A_159494).
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(6) M. Haverick, S. Mengisen, M. Shameem, and A. Ambrogelly, mAbs 6, 852–858 (2001).
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Pharm. Rev. 18, 59–63 (2015).(8) S. Fekete, A. Beck, and D. Guillarme,
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Biomedical Analysis 111, 169–176 (2015).(10) H.G. Barth and G.D. Saunders, LCGC
North Am. 30, 544–563 (2012). (11) P. Hong, S. Koza, and E.S.P. Bouvier, J.
Liq. Chrom. Rel. Techn. 35, 2923–2950 (2012).
(12) S. Fekete, A. Beck, J.L. Veuthey, and D. Guillarme, J. Pharm. Biomed. Anal. 101, 161–173 (2014).
(13) S. Fekete, K. Ganzler, and D. Guillarme, J. Pharm. Biomed. Anal. 78–79, 141–149 (2013).
(14) E. Gazal, Can size exclusion chromatography (SEC) be done on sub‑3‑μm particles?, presented at the 17th annual meeting of the Israel Analytical Chemistry Society, Tel Aviv, Israel (2014).
(15) S. M. Koza, P. Hong, K.J. Fountain, Advantages of ultra performance liquid chromatography using 125 Å pore size, sub‑2‑µm particles for the analysis of peptides and small proteins, poster presented at Medimmune, Rockville, MD, USA (2012).
(16) S. Fekete, J.L. Veuthey, and D. Guillarme, J. Pharm. Biomed. Anal. 69, 9–27 (2012).
(17) K. Sandra, I. Vandenheede, and P. Sandra, J. Chromatogr. A 1335, 81–103
(2014).(18) C. Horvath, B.A. Preiss, and S.R. Lipsky,
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Chromatogr. Sci. 7, 109–116 (1969).(20) S. Fekete, R. Berky, J. Fekete, J.L.
Veuthey, and D. Guillarme, J. Chromatogr.
A 1236, 177–188 (2012). (21) S.A. Schuster, B.M. Wagner, B.E. Boyes,
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Sep. Sci. 37, 189–197 (2014). (27) B. Bobály, D. Guillarme, and S. Fekete,
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(28) R. Hayes, P. Myers, T. Edge, H. Zhang, Analyst 139, 5674–5677 (2014).
(29) (A. Ahmed, W. Abdelmagid, H. Ritchie, P. Myers, and H. Zhankg, J. Chromatogr. A
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Szabolcs Fekete holds a PhD degree in analytical chemistry from the Technical University of Budapest, Hungary. He worked at the Chemical Works of Gedeon Richter Plc at the analytical R&D department for 10 years. Since 2011, he has worked at the University of Geneva in Switzerland. He has contributed 70 journal articles and authored book chapters. His main interests include liquid chromatography, column technology, pharmaceutical, and protein analysis. Jean-Luc Veuthey is professor at the School of Pharmaceutical Sciences, University of Geneva, Switzerland. He has also acted as President of the School of Pharmaceutical Sciences, Vice‑Dean of the Faculty of Sciences, and finally Vice‑Rector of the University of Geneva. His research domains include development of separation techniques in pharmaceutical sciences, and, more precisely, the study of the impact of sample preparation procedures in the analytical process; fundamental studies in liquid and supercritical chromatography; separation techniques coupled with mass spectrometry; and analysis of drugs and drugs of abuse in different matrices. He has published more than 300 articles in peer‑reviewed journals.Davy Guillarme holds a PhD degree in analytical chemistry from the University of Lyon, France. He is senior lecturer at the University of Geneva in Switzerland. He has authored 140 journal articles related to pharmaceutical analysis. His expertise includes HPLC, UHPLC, HILIC, LC–MS, SFC, and analysis of proteins and mAbs. He is an editorial advisory board member of several journals including Journal
of Chromatography A, Journal of
Separation Science, and LCGC
North America.
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Ph
oto
Cre
dit: Ju
no
s/G
ett
y Im
ag
es
It was Paul Ehrlich, who in around
1900, reported on “magic bullets” to
cure a wide range of diseases, thereby
indirectly referring to antibodies (1,2).
The development of the hybridoma
technology by Köhler and Milstein,
which allowed the production of
monoclonal antibodies (mAbs), bridged
the gap between concept and clinical
reality (3). Since the approval of the
first therapeutic murine mAb in 1986,
advances in antibody engineering has
allowed the production of chimeric
(mouse—human), humanized, and
human monoclonal antibodies, thereby
substantially improving safety and
efficacy and paving the way for the full
exploitation of mAbs for therapeutics
purposes (4,5). Over 40 mAbs are
now marketed in the United States and
Europe for the treatment of a variety
of diseases including cancer and
autoimmune diseases (6,7).
Eighteen displayed blockbuster status
in 2013 and six of these products had
sales of greater than $6 billion (Humira,
Remicade, Enbrel, Rituxan, Avastin,
and Herceptin). mAbs are currently
considered as the fastest growing class
of therapeutics with sales growing from
$39 billion in 2008 to almost $75 billion
in 2013, a 90% increase. Sales of other
recombinant protein biopharmaceuticals
have only increased by 26% in the same
time period, while small molecule drugs
are stagnating (6,7). The successes of
their predecessors have triggered the
development of various next-generation
mAb formats such as bispecific mAbs,
antibody–drug conjugates (ADC),
antibody mixtures, antibody fragments
(nanobodies, Fab), Fc fusion proteins,
and brain penetrant mAbs next to
glyco-engineered formats (4,5,8). With
several hundreds of products in (pre)
clinical development, the future looks
very bright.
The knowledge that the top-selling
mAbs are, or will become, open to
the market in the coming years has
resulted in an explosion of biosimilar
activities. Last year witnessed the
European approval of the first two
monoclonal antibody biosimilars
(Remsima and Inflectra), which both
contain the same active substance,
infliximab (9). Remicade, infliximab’s
blockbuster originator, reached global
sales of $8.9 billion in 2013. It is clear
that the biosimilar market holds great
potential but it is simultaneously
confronted with major hurdles. In
contrast to generic versions of small
molecules, exact copies of recombinant
mAbs cannot be produced because
of differences in the cell cloning and
the manufacturing processes used.
Even originator companies experience
lot-to-lot variability. As a consequence,
regulatory agencies evaluate biosimilars
based on their level of similarity to,
rather than the exact replication of, the
originator. In demonstrating similarity,
an enormous weight is placed on
analytics, and both biosimilar and
originator need to be characterized and
compared in great detail. In contrast to
small molecule drugs, mAbs are large
and heterogeneous (as a result of the
biosynthetic process and subsequent
manufacturing and storage), making
their analysis very challenging (10–13).
This article reports on selected state-
of-the-art chromatographic and mass
spectrometric (MS) tools for detailed
mAb characterization and comparability
assessment.
Protein A Chromatography for Clone Selection Protein A from Staphylococcus aureus
has a very strong affinity for the Fc
domain of IgG, allowing its capture from
complex matrices such as cell culture
supernatants. Affinity chromatography
making use of Protein A is the
gold standard in therapeutic mAb
purification and typically represents
the first chromatographic step in
downstream processing. Protein A
chromatography finds applications
beyond this large-scale purification. At
the analytical scale it is being used early
on in the development of mAbs for the
high-throughput determination of mAb
titre and yield directly from cell culture
supernatants and to purify µg amounts
of material for further measurements, for
Monoclonal Antibodies and Biosimilars — A Selection of Analytical Tools for Characterization and Comparability AssessmentKoen Sandra, Isabel Vandenheede, Emmie Dumont, and Pat Sandra, Research Institute for Chromatography (RIC) and
Metablys, Kortrijk, Belgium.
Monoclonal antibodies (mAbs) have emerged as important therapeutics for the treatment of life‑threatening diseases including cancer and autoimmune diseases. With the top‑selling mAbs evolving out of patent there has been a growing interest in the development of biosimilars. In demonstrating comparability to the originator product, biosimilar developers are confronted with an enormous analytical challenge. The article presents a selection of state‑of‑the‑art analytical tools for mAb characterization and comparability assessment.
16 Advances in Biopharmaceutical Analysis – October 2015
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example by mass spectrometry (MS) or
chromatography (14–16).
Figure 1 shows an overlay of the
Protein A chromatograms of 12
trastuzumab-producing Chinese
Hamster Ovarian (CHO) clones,
generated in the framework of a
Herceptin biosimilar development
programme. Herceptin (scientific INN
name trastuzumab) is being used in
the treatment of HER2 positive breast
cancer. It is open to the European
market and evolves out of patent in
the US in 2019 (17). Given its market
potential (global sales of $6.5 billion in
2013), dozens of companies are actively
developing a Herceptin biosimilar. The
unbound CHO material elutes in the
flow-through while the mAb is captured
and only released after lowering the
pH. From these chromatograms,
a distinction can already be made
between low and high mAb producing
clones. Absolute mAb concentrations
can be determined by linking the peak
areas to an external calibration curve
constructed by diluting Herceptin
originators. Obtained mAb titres are
visualized in the bar plot in Figure 1.
From the findings, clear decisions
can be made for further biosimilar
development, that is, high trastuzumab
producing clones can be selected and
taken further in development.
Next to the mAb titre, the second
important criterion in clone selection
is based on the structural aspects. In
the case of biosimilar development, the
structure should be highly similar to the
originator product, within the originator
batch-to-batch variations. Figure 2
shows the ion mobility (IM) quadrupole
time-of-flight (QTOF) MS measurements
of inter-chain reduced Herceptin and
Protein A purified trastuzumab from a
high titre CHO clone. Samples were
introduced into the MS system via
a reversed-phase on-line desalting
cartridge and light (Lc) and heavy
chain (Hc) were resolved in the IM drift
cell. Two Lc forms with identical m/z
and molecular weight (MW) but with a
different drift time, hence conformation,
are highlighted. Deconvoluted spectra
reveal that clone derived trastuzumab
and originator display the same Lc
and Hc MW values. In addition, the
same N-glycans, which are of the
complex type, are observed on the
Hc of the originator and clone derived
mAb. These are considered the most
important attributes of biosimilarity
according to US and European
regulatory authorities (primary sequence
should be identical and glycosylation
should be preserved). While
glycosylation is similar from a qualitative
perspective, quantitative differences
are observed. Our experience in clone
selection studies has found that it is
not always the case that MW values
are identical between originators and
mAbs derived from high titre clones
or subclones. In these situations,
mAbs are typically not taken further in
development.
Reversed‑Phase LC–MS Analysis of Intact, Reduced, Papain, and IdeZ Cleaved mAbWhen a mAb is taken further in
development, a detailed characterization
and comparability assessment
has to be performed. Structural
characteristics such as amino acid
sequence and composition, molecular
weight and structural integrity, N- and
O-glycosylation, N- and C-terminal
17www.chromatographyonline.com
Sandra et al.
0,8
0,7
0,6
0,5
0,4
0,3
0,2
0,1
0,03
Co
nce
ntr
ati
on
(m
g/m
L)
6 8 9 10 14 24
Clone
Time (min)
mA
U
21.510.50
700
600
500
400
300
200
100
25 26 27 28 32
0
mAb
Figure 1: Overlaid UV 280 nm Protein A chromatograms of 12 trastuzumab producing CHO clones with graphical representation of the mAb titre, expressed in mg/mL.
mAb structure: mAbs are tetrameric immunoglobulin G (IgG) molecules with
a MW of 150 kDa composed of two light (Lc – 25 kDa) and two heavy (Hc – 50
kDa) polypeptide chains connected through inter-chain disulphide bridges.
Twelve intra-chain disulphide bridges, four within each Hc and two within
each Lc, furthermore guarantee its structural integrity. Six different globular
domains, that is, one variable (VL) and one constant domain (CL) for the Lc
and one variable (VH) and three constant domains (CH1, CH2, CH3) for the
Hc, are recognized. The structure can also be divided in the antigen-binding
fragment (Fab), composed of VL, CL, VH, and CH1 and the crystallizable
fragment (Fc) composed of CH2 and CH3. Antigen-binding is mediated
by the Fab fragment while the Fc fragment is responsible for the effector
function, that is, antibody-dependent cell-mediated cytotoxicity (ADCC) and
complement-dependent cytotoxicity (CDC). All mAbs are glycoproteins with
two conserved N-glycosylation sites in the Fc region that can be occupied
with complex and high mannose type N-glycans. These glycan structures are
known to play a role, amongst others, in the effector function.
ES683294_LCESUPP1015_017.pgs 10.01.2015 20:49 ADV blackyellowmagentacyan
processing, S-S bridges, deamidation
(asparagine, glutamine), aspartate
isomerization, and oxidation (methionine,
tryptophan) need to be assessed. In that
respect, reversed-phase LC is extremely
powerful. Figure 3 shows highly efficient
reversed-phase LC–UV chromatograms
obtained on intact, inter-chain reduced,
papain digested, and non-reduced and
reduced IdeZ cleaved Herceptin. All
these chromatograms are generated
using exactly the same chromatographic
conditions making use of widepore
sub-2-µm C8 particles, elevated
column temperatures (80 °C) and
trifluoroacetic acid as ion-pairing
reagent in a water/acetonitrile mobile
phase system. Under these conditions,
many of the challenges encountered
in performing reversed-phase LC of
proteins (peak tailing, peak broadening,
and adsorption) are tackled (18–19).
Moreover, these conditions are
compatible with MS, which allows
an in-depth characterization and
comparability assessment of mAbs.
Figure 4 shows the reversed-
phase LC–UV–MS analysis of IdeZ
cleaved and TCEP reduced Herceptin
originator and biosimilar. IdeZ or
immunoglobulin-degrading enzyme from
Streptococcus equi ssp zooepidemicus
is a highly specific protease similar to
IdeS that cleaves mAbs at a single site
below the hinge region, yielding F(ab’)2
and Fc/2 fragments (20, 21). Following
reduction, the F(ab’)2 fragment is
converted into the Lc and Fd’. From the
simultaneously acquired MS data it can
18 Advances in Biopharmaceutical Analysis – October 2015
Sandra et al.
40
35
30
251000 1500
Drift Time (ms) vs. m/z
2000 2500 1290 1295 1300 13101305
Drift Time (ms) vs. m/z
1315
195025
30
35
40
32
31
34
33
36
35
38
37
1952 1954 1956 1958Drift Time (ms) vs. m/z
1960
Heavy chain
44+
12+
12+ 11+10+
9+
13+
13+
14+15+
16+17+
18+
40+38+
36+
42+
Light chain
Light chain
Light chain (12+)
Light chain (12+)
Light chain 1 (18+)
G0G0F G1F
G2F
Heavy chain (39+)
23100 23200 23300
23439.9
23440.1
0
1
2
3
x106
0
1
2
3
x106
0
1
2
3
4
x106
0
1
2
x106
Counts vs. Deconvoluted Mass (amu) Counts vs. Deconvoluted Mass (amu)
23400 23500 23600 23700 23800 23900 49800 50200
50598.4
50760.6
50922.4
50922.850452.3
50598.150760.7G0F
G0 G2F
Originator
Biosimilar
Originator
Biosimilar
G1F
50600 51000 51400 51800
Figure 2: IM-QTOF-MS profile of inter-chain reduced Herceptin (top). Deconvoluted light and heavy chain spectra of a Herceptin originator and a trastuzumab-producing clone (bottom).
Reduction
Reduction
Papain
2 * Lc 2 * Hc
2 * Fab 2 * Fc
1 * F(ab)’2
F(ab)’2 Fd’
Fc/2
Lc
Fc
Fab
Hc
Lc
2 * Fc/2 2 * Lc 2 * Fd’ 2 * Fc/2
Fc/2
IdeZ
6 8 10 12 14 16 18
6 8 10 12 14 16 18
20
64 8 10 12 14 16 18 20 228 10 12 14 16 18 20 22
Time (min)
Time (min)
Time (min)
Time (min)
Time (min)
10 12 14 16 18 20 22
mA
U
mA
U
mA
Um
AU
mA
U
200
150
100
50
0
500
400
300
200
100
0
500
400
300
200
100
0
500
400
300
200
100
0
240
200
160
120
80
0
Figure 3: Reversed-phase LC–UV separations of intact, dithiothreitol (DTT) reduced, papain digested, non-reduced IdeZ cleaved and tris(2-carboxyethyl)phosphine (TCEP) reduced IdeZ cleaved Herceptin. These represent extremely powerful separations for comparability assessment and for detailed characterization. Conditions are compatible with MS allowing identification of the observed peaks.
ES683297_LCESUPP1015_018.pgs 10.01.2015 20:49 ADV blackyellowmagentacyan
be deduced that peaks b, b’, d, d’, g,
and g’, corresponding to, respectively,
Fc/2, Lc and Fd’, are identical in both
the originator and biosimilar. The
measured MW values obtained are
well below 0.005% of the theoretical
MW values, which is expected when
using high-resolution and accurate
mass instrumentation. Upon examining
the spectra of the Fc/2 fragment, the
biosimilar appears to be enriched in
the N-glycan G0F while a more even
distribution between G0F and G1F is
observed in the originator. This is also
reflected in the chromatographic peak
shape. The broader peak b’ indicates
a partial separation of the G0F and
G1F species, with the former eluting
slightly later. Several other differentiating
peaks are observed in the separation
of the biosimilar, that is, peaks a, c, e,
and f. Compared to peak b, peak a
displays a 128 Da mass increase, which
can be explained by the presence of
a C-terminal lysine. To provide some
more background on this particular
event, the Hc is cloned with a lysine
residue at the C-terminus. During protein
maturation, this lysine is removed by
19www.chromatographyonline.com
Sandra et al.
Fc/2
Lc Fd’
Biosimilar
Originator
b
a
b’
8 9 10 11 12 13
Response Units vs. Acquisition Time (min)
x10 2
x10 2
2
1.5
1
0.5
2.5
x10 3
x10 4
x10 4
5
x10 3
8
6
4
2
0
x10 4
8
x10 5
1.25
1
0.75
0.5
0.25
0
6
4
2
0
Fc/2 + K(G0F) a
b
b’
25365.2
Fd’ + 1 Hex
Fd’ + 2 Hex
f
g
g’
25707.6
Fd’
25384.0
Fd’25384.3
25546.1
x10 3
x10 4
x10 3
x10 5
1
0.5
0
4
2
S
S
S
S
S
N
DS
S
S
S
0
2
3
2
1
0
0
7.5
5
2.5
Lc + 2 Hex
Lc + 1 Hex
Lcd
c
d’
e
23766.8
23605.4
23443.8
Lc + deam23444.7
Lc23443.7
Fc/2 (G0F)
NHYTQKSLSLSPG
NHYTQKSLSLSPGK
25237.0
Fc/2 (G1F)25399.2
Fc/2 (G1F)
Fc/2 (G2F)
25399.1Fc/2 (G0F)
Fc/2 (G0)
25236.9
25090.7
25560.9
25254.0
4
3
2
1
0
4
3
2
1
0
2.5
2
1.5
1
0.5
024800 24900 25000 25100 25200 25300 25400
Counts vs. DeconvolutedMass (amu)
24800 25000 25200 25400 25600 25800 26000 26200 26400 2660024600
Counts vs. DeconvolutedMass (amu)
22800 23000 23200 23400 23600 23800 24000 24200 24400
Counts vs. DeconvolutedMass (amu)
25500 25600 25700 25800 25900 26000
2
1.5
1
0.5
0
0
14 15 16 17 18 19 20 21
d’g’
c
d
ef
g
Figure 4: Reversed-phase LC–UV–MS analysis of IdeZ cleaved and TCEP reduced Herceptin originator and biosimilar and deconvoluted MS spectra associated with the annotated peaks.
T45 + G2F
Originator
Originator
Biosimilar
Biosimilar
Originator
Biosimilar
Originator
Biosimilar
T45 + G0F
T45 + G1Fb
T45 + G1FaT45 + G0
T45 + G0F
T45 + G1F T45 + G0
T62
T62
T3
T3
13.1%T3 deam
10.5%T62+K
0
20
40
60
80
100
120
0
20
40
60
80
5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33
10.6 10.8 11 11.2 15.4 16.2 17 17.8 18.4 18.8 19.2
Response Units vs. Acquisition Time (min)
Figure 5: Reversed-phase LC–UV peptide map of Herceptin originator and biosimilar with detail in some specific regions showing post-translational modifications. T45: EEQYNSTYR, T62: SLSLSPG, T3: ASQDVNTAVAWYQQK. Peak identities were assigned by the simultaneously acquired MS and MS–MS data.
ES683296_LCESUPP1015_019.pgs 10.01.2015 20:49 ADV blackyellowmagentacyan
host cell carboxypeptidases. This
process is more dominant in the host
cell producing the originator product
than in the host cell producing the
biosimilar mAb. From the MS data it can
be deduced that peak c originates from
the Lc plus 1 and 2 hexose units. This
potentially originates from a glycation
event, which appears to be negligible
in the originator mAb. Peak e shows a 1
Da mass increase compared to peak d,
indicating a deamidation in the Lc. This
event is apparent in both the originator
and biosimilar with an increased
occurrence in the biosimilar. In analogy
with peak c, peak f displays 162 Da
spacings on Fd’, which is indicative of
glycation.
Reversed‑Phase LC–MS Analysis for Peptide MappingAs previously demonstrated, protein
measurement is extremely powerful
but does not provide the complete
picture. While it is indicative for identity
and highlights dominant modifications,
it does not provide the actual amino
acid sequence nor does it localize
the modifications. For example, the
measurement presented in Figure 4
reveals a deamidation on the Lc (peak
e) but it cannot be traced back to
a specific asparagine or glutamine
residue. The Lc of the measured
mAb contains six asparagine and 15
glutamine residues, which are all prone
to this chemical modification. These
characteristics can further be assessed
at the peptide level following proteolytic
digestion. When digesting Herceptin
with the enzyme trypsin, which cleaves
the protein next to arginine and
lysine residues, 62 identity peptides
are formed. Taking into account
post-translational modifications and
incomplete and aspecific cleavages
taking place, over 100 peptides with
varying physicochemical properties
in a wide dynamic concentration can
be expected. This is a particularly
complex sample and demands the
best in terms of separation technique.
Again, reversed-phase LC is the
method of choice to resolve these
complex mixtures. Figure 5 shows the
UV peptide maps of both the originator
and biosimilar. By taking advantage of
the simultaneously acquired MS data,
over 99% of the peptide sequence
can be covered in both the originator
and biosimilar thereby confirming
identity. While peptide maps are
highly comparable, differences in
post-translational modifications can be
detected (Figure 5). Obtaining a good
knowledge of all of these modifications
is important since they could be
critical to the potency and safety of a
mAb. A deviating glycosylation profile
between originator and biosimilar is
already revealed at the protein level
(Figure 4). At the peptide level, the
different N-glycosylated variants are
nicely resolved chromatographically
and are shown to be located on
peptide EEQYNSTYR. Again, the
undergalactosylation of the biosimilar
is apparent. The peptide map also
reveals the presence of a lysine at the
C-terminal peptide of the heavy chain
(SLSLSPGK) and slightly increased
deamidation in a light chain peptide
(ASQDVNTAVAWYQQK). This particular
peptide contains four potential
deamidation sites (3 Gln and 1 Asn).
Based on MS measurement one cannot
discriminate between the four sites.
Upon performing MS–MS and carefully
interpreting the fragment ions observed,
the deamidation can be traced back
to the N (11). This deamidation in fact
corresponds to the deamidation event
20 Advances in Biopharmaceutical Analysis – October 2015
Sandra et al.
CEX
SEC
HIC
Asndeamidation
mA
Um
AU
mA
U
1000
800
600
400
200
35
350
300
250
200
150
100
50
0
30
25
20
15
10
5
0
2
10 12 14 16 18 20
4 6 8 10 12 14 16
0
12.5 15 17.5 20 22.5 25 27.5 30 32.5
Buffer excipients
Dimer0.4%
Time (min)
Time (min)
Time (min)
Figure 6: CEX, SEC, and HIC separations of Herceptin. These techniques are used in characterization and release testing.
ES683298_LCESUPP1015_020.pgs 10.01.2015 20:49 ADV blackyellowmagentacyan
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ES683548_LCESUPP1015_021_FP.pgs 10.01.2015 21:53 ADV blackyellowmagentacyan
observed in the reduced IdeS digest
(Figure 4). At that time this event could
be linked to the Lc but could not be
traced back to a specific residue.
As discussed, mAb digests can
be quite complex and their analysis
demands the best in terms of separating
power. If one-dimensional (1D)
separations are not able to provide
the separation power needed, one
can opt for two-dimensional (2D)
LC. Compared to 1D-LC, 2D-LC and
especially comprehensive LC (LC × LC)
will drastically increase resolution. We
have recently described the analysis
of Herceptin originator and biosimilar
digests on the combination reversed-
phase LC × reversed-phase LC (22). It is
important to point out that orthogonality
in reversed-phase LC × reversed-phase
LC peptide mapping is only obtained
when operating the two reversed-phase
LC columns at different pH values.
This is a direct result of the zwitterionic
nature of peptides, which gives rise
to major selectivity differences at pH
extremes. These reversed-phase LC
× reversed-phase LC peptide maps
provide a wealth of information and allow
both identity and purity to be assessed.
This makes it an attractive technology for
the comparison of different production
batches and to compare innovator
biopharmaceuticals with biosimilars.
Native Chromatographic Tools: Size‑Exclusion Chromatography, Cation‑Exchange Chromatography, and Hydrophobic Interaction ChromatographyIn contrast to reversed-phase LC,
size-exclusion chromatography
(SEC), ion-exchange chromatography
(IEX), and hydrophobic interaction
chromatography (HIC) are
non-denaturing techniques that provide
complementary information to the
afore-mentioned chromatographic
mode (Figure 6). These techniques are
used early on in mAb characterization
and comparability assessment and
are subsequently applied in routine
testing. A major advantage of these
chromatographic modes over
reversed-phase LC is that they preserve
the structure, and so minor variants
can be collected and subjected to
complementary techniques such as
potency determination. SEC, IEX,
and HIC are not directly compatible
with MS because of the presence of
non-volatile salts in the mobile phases.
The identification of peaks requires their
collection and subsequent desalting
or dilution prior to MS measurement.
Desalting of the collected fractions
can be performed in an automated
manner using, for example, a small
reversed-phase cartridge hyphenated to
an MS system.
Ion-exchange chromatography
is an excellent tool to highlight
charged variants that might
arise from modifications such as
deamidation, lysine truncation, or
N-terminal cyclization (23,24). Since
most therapeutic mAbs have a
higher proportion of basic residues,
cation-exchange chromatography (CEX)
is the most commonly used technique.
The CEX separation of Herceptin
(Figure 6) highlights the asparagine
deamidation discussed earlier. A
deamidation renders a protein more
acidic, which explains this earlier elution.
Size-exclusion chromatography is
the chromatographic mode with the
lowest efficiency or resolution of the
afore-mentioned techniques, but it is
extremely powerful when determining
aggregation and fragmentation. It
is recognized that aggregates may
stimulate immune responses and it is
therefore very important to measure this
critical quality attribute. Aggregation
can typically not be assessed using
22 Advances in Biopharmaceutical Analysis – October 2015
Sandra et al.
(a) OriginatorG0F
G0F
G1Fa
G1FaMan5G0F-G
lcN
Ac
G1Fb
G1Fb
G2FG0
Biosimilar
LU14
12
10
8
6
4
2
0
LU
14
16
10 12.5 15 17.5 20 22.5 25 27.5
(b) Biosimilar
Biosimilar: 4x
Biosimilar: 8x
Biosimilar: 16x
Biosimilar: 24x
LU
10
10
5
0
LU
8
4
0
LU
8
4
0
LU8
4
0
LU8
4
0
12.5 15 17.5 20 22.5 25 27.5
10 12.5 15 17.5 20 22.5 25 27.5
10 12.5 15 17.5 20 22.5 25 27.5
10 12.5 15 17.5 20 22.5 25 27.5
10 12.5 15 17.5 20 22.5 25 27.5
Time (min)
Time (min)
Time (min)
Time (min)
Time (min)
Time (min)
Time (min)
10 12.5 15 17.5 20 22.5 25 27.5
12
10
8
6
4
2
0
Figure 7: (a) Overlaid HILIC-FLD chromatograms of the 2-AB labelled N-glycans enzymatically released from Herceptin originator and Protein A purified biosimilar. (b) N-glycan profiles of the biosimilar obtained by growing the CHO clone at different galactose, uridine, and manganese chloride concentrations. Separations were performed on superficially porous HILIC particles.
ES683295_LCESUPP1015_022.pgs 10.01.2015 20:49 ADV blackyellowmagentacyan
the other chromatographic modes
discussed. Figure 6 presents the SEC
analysis of Herceptin and illustrates that
dimers can be measured accurately at
levels as low as 0.4%.
In recent years, HIC has been
revisited mainly from the perspective
of ADC’s governing a separation
based on the number of conjugated
drugs allowing the drug-to-antibody
ratio (DAR) to be determined. In the
separation of naked mAbs it is useful
to highlight heterogeneities originating
from oxidation, aspartate isomerization,
deamidation, succinimide formation,
C-terminal lysine, and clipping. The
HIC analysis of Herceptin gives rise to a
single chromatographic peak (Figure 6).
Hydrophilic Interaction Liquid Chromatography for Glycan ProflingAs demonstrated, glycosylation can
be revealed at both the protein and
peptide level. A detailed insight into
the sugars, however, can only be
obtained following their removal from
the protein/peptide backbone. This is
preferably done enzymatically using the
deglycosidase PNGase F. The liberated
sugars are subsequently labelled
via reductive amination to improve
their chromatographic separation
and detectability (fluorescence and/
or mass spectrometric detection).
The fluorescence trace is typically
used for quantitative purposes while
the MS trace is used for qualitative
purposes. Figure 7(a) displays the
analysis of 2-aminobenzamide (2-AB)
labelled Herceptin originator and
biosimilar N-glycans using hydrophilic
interaction chromatography (HILIC)
with fluorescence detection (FLD).
In this particular case, a column
packed with superficially porous HILIC
particles compatible with 600 bar
HPLC instrumentation was used. This
measurement provides information
on the glycans and allows structural
isomers, that is, G1Fa and G1Fb which
differ in the positioning of the galactose
residue either on the α1-3 or α1-6
branch of the complex type glycan, to
be resolved.
The same type of complex N-glycans
are observed on both the originator and
biosimilar but quantitative differences
are revealed with an overexpression of
G0F species on the biosimilar, which is
in accordance with the measurements
performed at protein and peptide level.
Since glycosylation is a critical quality
attribute, this undergalactosylation does
not make the product similar enough to
be considered by regulatory authorities
as a Herceptin biosimilar.
The biosimilar-producing CHO cell
culture medium was subsequently
tuned by feeding uridine (U), galactose
(G), and manganese chloride (M) at
different concentrations (25). These
are the substrates and activator of the
galactosyltransferase responsible for
donating galactose residues to G0F and
G1F acceptors. Figure 7(b) shows the
N-glycan profiles obtained by growing
the biosimilar producing CHO clone at
different U, G, and M concentrations.
It is observed that the ratio G1F/G0F
increases with increasing concentration
of U, G, and M. From these results it
can be concluded that conditions can
be found that allow the glycosylation of
the biosimilar to fit within the originator
specifications.
ConclusionIn the development of biosimilars, a
comprehensive comparability exercise
involving the originator product is
required to demonstrate similarity in
terms of physicochemical characteristics,
efficacy, and safety. In that respect,
an enormous weight is placed on
analytics and the analytical package
for a biosimilar mAb submission is
considerably larger than that of a
stand-alone mAb. Structural differences
define the amount of pre-clinical and
clinical studies required. A wide range of
analytical tools providing complimentary
information is available to guide biosimilar
development.
AcknowledgementsThe authors acknowledge Maureen
Joseph (Agilent Technologies,
Wilmington, USA), David Wong (Agilent
Technologies, Santa Clara, USA) and
Lindsay Mesure (Promega, Leiden, The
Netherlands).
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Anal. Biochem., 415, 212–214 (2011).
(21) C. Hosfield, P. Compton, L. Fornelli, P.
Thomas, N.L. Kelleher, M. Rosenblatt,
and M. Urh, Promega Poster Part#PS260
(2015).
(22) G. Vanhoenacker, I. Vandenheede, F.
David, P. Sandra, and K. Sandra, Anal.
Bioanal. Chem. 407, 355–366 (2015).
(23) I. Vandenheede, E. Dumont, P. Sandra, K.
Sandra, M. Joseph, Agilent Technologies
Application Note 5991-5273EN (2014).
(24) I. Vandenheede, E. Dumont, P. Sandra, K.
Sandra, M. Joseph, Agilent Technologies
Application Note 5991-5274EN (2014).
(25) M.J. Gramer, J.J Eckblad, R. Donahue,
J. Brown, C. Schultz, K. Vickerman, P.
Priem, E.T. van den Bremern J. Gerritsen,
and P.H. van Berkel, Biotechnol. Bioeng.
108, 1591–1602 (2011).
Koen Sandra is Director at the
Research Institute for Chromatography
(RIC, Kortrijk, Belgium).
Isabel Vandenheede is a Protein
Analyst at the Research Institute
for Chromatography (RIC, Kortrijk,
Belgium).
Emmie Dumont is an LC–MS
Specialist at the Research Institute
for Chromatography (RIC, Kortrijk,
Belgium).
Pat Sandra is Chairman at the
Research Institute for Chromatography
(RIC, Kortrijk, Belgium) and Emeritus
Professor at Ghent University (Ghent,
Belgium).
23www.chromatographyonline.com
Sandra et al.
ES683299_LCESUPP1015_023.pgs 10.01.2015 20:49 ADV blackmagentacyan
Monoclonal antibodies (mAbs) and
their related products are the fastest
growing class of human therapeutics
(1). Sixty IgGs and derivatives
(antibody drug conjugates [ADCs],
radio-immunoconjugates, bispecific
antibodies, Fab fragments, and
Fc-fusion proteins and peptides)
have been approved for use in
various arenas such as cancers,
inflammatory diseases, and, more
recently, for the treatment of high
cholesterol. In oncology, however,
the first generation mAbs often
lack efficiency or face resistance
as a result of upregulation of HER2
downstream signalling pathways
in the case of trastuzumab for
example (1). ADCs are emerging
as an important subclass of armed
immunoglobulins (2), with two
approved first-in-class drugs, namely
brentuximab vedotin (Adcetris,
Seattle Genetics/Takeda) and
trastuzumab emtansine (Kadcyla,
Genentech/Roche), now on the
market. Importantly, 50 more ADCs
are now investigated in clinical trials
in many different types of cancers.
They are the result of numerous
technological improvements
based on a better understanding
of structure-function relationships,
thanks in no small part to state-of-
the-art mass spectrometry (MS),
which will be discussed below (4).
The Anatomy of Antibody Drug ConjugatesADCs (around 154 kDa) are
constructed from three components:
a mAb (around 148 kDa) that is
specific to a tumour antigen, a
highly potent cytotoxic agent, and a
chemical linker (0.3 to 1.5 kDa) that
enables covalent attachment of an
average of four cytotoxic payloads
to the mAb (Figure 1). For most
ADCs, the primary sites used for
protein-directed conjugation are the
sulphydryl groups of the inter-chain
cysteine residues or amino groups
of lysine residues of the mAbs (5).
Alternatively, glycans or engineered
amino acids or tags can be used
as attachment sites to yield
more homogeneous site-specific
conjugates, which often results
in an improved therapeutic
index (6).
The ADC Analytical ToolboxAs highlighted in Figure 2, a
combination of native and denaturing
methods are mandatory to gain
structural insights of IgG hinge
cystein-linked drug conjugates.
In analogy to brentuximab
vedotin, nearly two-thirds of the
immunoconjugates currently in
clinical trials are produced through
partial reduction of the four
inter-chain disulphide bonds of IgG1
antibodies (chimeric, humanized, or
human), followed by alkylation with
a preformed drug-linker maleimide
activated species (Figure 3[a]).
This process results in conjugates
with a distribution of 0, 2, 4, 6 or 8,
drugs incorporated per antibody and
an average drug to antibody ratio
(DAR) of 4 drugs/mAb (7). This is
routinely controlled by hydrophobic
interaction chromatography (HIC)
under non denaturing conditions
(8) (Figure 3[b]). In an orthogonal
way, these structures are confirmed
both by reversed-phase high
performance liquid chromatography
(HPLC) under reducing conditions
and capillary electrophoresis sodium
dodecyl sulphate (CE–SDS) under
both non-reducing and reducing
conditions (9). Ultimately, MS
methods are used for structural
assessment (10); this includes
hydrogen-deuterium exchange
MS (HDX-MS) for interrogating
the higher-order structure of
ADCs, as recently reported by
Valliere-Douglass et al. (11).
For the mAb moiety, dozens of
Harnessing the Benefits of Mass Spectrometry for In-depth Antibody Drug Conjugates Analytical CharacterizationAlain Beck1 and Sarah Cianférani2,3, 1Centre d’Immunologie Pierre-Fabre (CIPF), Saint-Julien-en-Genevois, France, 2BioOrganic Mass Spectrometry Laboratory (LSMBO), IPHC, Université de Strasbourg, Strasbourg, France, 3IPHC, CNRS,
UMR7178, Strasbourg, France.
Antibody drug conjugates (ADCs) are a fast growing class of empowered anti-cancer biopharmaceuticals. Also known as immunoconjugates, ADCs are composed of cytotoxic drugs covalently attached via a conditionally stable linker to monoclonal antibodies (mAbs) highly specific for tumour-associated antigens. Compared to naked mAbs, ADCs have an increased level of complexity as the heterogeneity of conjugation cumulates with the inherent microvariability of the immunoglobulins (IgGs). This article highlights recent progress in high-resolution mass spectrometry (HRMS) and liquid chromatography–mass spectrometry (LC–MS) methods for the structural characterization of brentuximab vedotin and trastuzumab emtansine, two FDA and EMA approved ADCs directed against haematological and solid tumours, respectively.
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Advances in Biopharmaceutical Analysis – October 201524
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microvariants have been identified
and reported in the literature such as
glycoforms, charge, cysteine-related,
oxidized, size, and low level point
mutation variants (12). Conjugation
of payloads to mAbs increases the
structural complexity of the resulting
molecule (13), which triggers the
need for improved characterization
methods for the analysis of drug
loading and distribution, average
DAR, size and charge variants,
un-conjugated drug-linker, and ADC
biophysical properties (14).
As for antibodies, ADCs are
analyzed at different levels (top,
middle, and bottom) as illustrated
below for brentuximab vedotin
and trastuzumab emtansine, two
first-in-class and gold standards
for cysteine and lysine conjugated
ADCs, respectively.
Brentuximab Vedotin Characterization Under Native Conditions (Top Level, 148 to 160 kDa)Using native desalting conditions,
Valliere-Douglass et al. reported
the expected mass measurement
of the intact bivalent structure
of the hinge-cysteine linked
ADCs (15), which would normally
decompose as a consequence of
the denaturing chromatographic
conditions typically used for
liquid chromatography−mass
spectrometry (LC−MS). The mass of
the desalted ADC was determined
using standard desolvation and
ionization conditions. Successful
intact mass measurement of
IgG1 mAbs conjugated with
maleimidocaproy-monomethyl
Auristatin F (mcMMAF) and
valine-citrulline-monomethyl
Auristatin E (vcMMAE) at inter-chain
cysteine residues were reported. This
method was also used to detect the
changing drug load distribution over
time from a set of in vivo samples
(16). As an alternative, Chen et al.
reported the use of limited enzymatic
digestion with a cysteine protease for
vcMMAE characterization by native
MS with an improvement of low
abundance D6 and D8 species (17).
Tandem native MS on brentuximab
vedotin was successfully used
by Dyachenko et al. to show that
drug conjugation takes place non
homogeneously to cysteine residues
both on the light and heavy chains
(18).
In addition, Debaene et al. recently
reported on the combination of
native MS to ion mobility MS (IM–MS)
for ADC characterization (19). As a
proof of principle, they highlighted
the benefits of high-resolution native
MS (Figure 3[c]) and native IM–MS
for the determination of the drug
load profile, naked antibody content,
and average DAR of brentuximab
vedotin (Figure 4). The analytical
potential of native MS and IM–MS
was compared to HIC, the gold
standard for ADC quality control.
The benefits of high-resolution
native MS were demonstrated for
drug distribution and average DAR
determination along with improved
mass accuracies (<30 ppm in routine
analysis). The main advantage of
using native MS for exact mass
measurements of ADCs with
inter-chain cysteinyl-linked drugs lies
in its ability to detect non-covalent
associations of light and heavy
chains that cannot be directly
analyzed by classic denaturing
LC–MS methods. Interestingly,
heterogeneity of drug loading on
mAbs was uniquely evidenced by
differences in drift times. Collisional
cross sections were measured for
each payload species and affirmed
slight conformational changes
induced by drug conjugation. Finally,
a semi-quantitative interpretation
of IM–MS data was presented
that allowed the average DAR and
DAR distribution to be directly
extrapolated. Both native MS
and IM–MS experiments were in
agreement with results obtained
from HIC. For full proof of principle,
Advances in Biopharmaceutical Analysis – October 201526
Beck and Cianférani
(a) Drug-linker300–1500 Da
(b) mAb148,000 Da
(c) ADC (avDAR 4)154,000 Da (+ 4%)
Figure 1: 3D-models of (a) drug-linkers, (b) mAbs, and (c) ADCs.
Drug loadprofle
AverageDAR
Un-conjugatedmAb (D0)
Free drug-linker (DL)
and relatedimpurities
Higher orderstructures
(HOS) Conjugation sitesand positional
isomers
Denaturing methods• nr/rSDS-PAGE
• nr/rCE-SDS
• LC–MS (+/- Red ; IdeS)
• Peptide mapping
Native methods• UV
• HIC
• SEC
• A4F
• Native MS
• Ion mobility MS
Figure 2: The ADC analytical toolbox: a panel of orthogonal separation and structural methods in native and denaturing conditions.
ES683363_LCESUPP1015_026.pgs 10.01.2015 21:02 ADV blackyellowmagentacyan
HIC fractions were collected and
analyzed by native MS and IM–MS,
allowing HIC limitations, for example
in the case of highly hydrophobic
payloads, to be circumvented
(Figure 4).
Brentuximab Vedotin Subunit Characterization under Denaturing and Reducing Conditions (Middle Level, 23 to 54 kDa)For hinge cysteine-conjugated
ADCs, most of the inter-chain
disulphide bridges are no longer
present but are replaced with the
linker-drugs during conjugation
and the ADC is held together
through non-covalent hydrophobic
interactions (Figure 3[a]).
Reversed-phase HPLC offers an
orthogonal method for drug load
profiling and for calculating the
average DAR. In analogy to HIC,
reversed-phase HPLC is also based
on hydrophobicity differences but
the use of organic solvent and
a small amount of organic acid
instead of salt is disruptive for the
intact cysteine-linked ADC. When
analyzed directly, the ADC cannot
withstand the highly denaturing
conditions and will dissociate into
antibody fragments. Treatment
of the cysteine-linked ADC with
dithiothreitol (DTT) or (tris(2-
carboxyethyl)phosphine) (TCEP)
fully reduces the remaining inter-
chain disulphides and yields six
species: light chain with 0 and 1
drug attached (L0 and L1), and
heavy chain with 0, 1, 2, or 3 drugs
attached (H0, H1, H2, and H3).
These species are stable in the
denaturing organic environment
and can be well resolved on a
reversed-phase column such as
PLRP-S. The weighted average DAR
is obtained by integration of the
light and heavy chain peaks and
calculation of the percentage peak
area; the assigned drug load for
each peak must also be taken into
account (20).
Brentuximab Vedotin Subunit Characterization under Denaturing Conditions after IdeS Digestion and Reduction (Middle Level, 23 to 28 kDa)The immunoglobulin-degrading
enzyme of Streptococcus pyogenes
(IdeS, Fabricator, Genovis) is
becoming increasingly popular
for the fast characterization of
antibodies by MS, including correct
sequence assessment, antibody
Fab and Fc glyco-profiling,
biosimilar comparability studies,
and Fc-fusion protein studies. IdeS
specifically cleaves immunoglobulin
G and related products under
27www.chromatographyonline.com
Beck and Cianférani
3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 11.00
D0
D2 D4
D6
D8
(c) Native mass spectrometry
(b) HIC profle
MaxEnt deconvolution
146
Mass (kDa)
Time (min)
150 148 154 152 158 156
D4
D2
D6
D8 D0
1. Reduction 2. Conjugation
+2635.1 Da
+2634.3 Da +2635.4 Da
+2634.2 Da
(a) Mild reduction and conjugation process
8 interchaincysteines
DAR = 3.9
= 4.0 DAR
DAR
8nA
n0Σ
DAR
8A
n0Σ
Figure 3: Hinge Cys conjugates: (a) conjugation process, (b) native chromatographic (HIC), and (c) mass spectrometry methods (ESI–MS).
(0.08_1.96) (1.00_200.00) 1000.00_7998.00) 023564FDE.raw : 1 max: 6993 m/z
6500
m/z
6000
• 6500
• 6000
• HIC D0 • HIC D2 • HIC D4 • HIC D6 • HIC D8
12 14 16Drift Time (milli secs)
18 20
Figure 4: Native IM–MS: brentuximab vedotin driftscope plots (drug substance and individual HIC separated D0, D2, D4, D6, and D8 peaks).
ES683365_LCESUPP1015_027.pgs 10.01.2015 21:02 ADV blackyellowmagentacyan
its hinge domain. As shown by
Wagner-Rousset et al., IdeS is
also a powerful enzyme for fast
characterization (drug loading,
distribution, and average DAR) of
ADCs and antibody-fluorophore
conjugates (AFC). It also allows
glyco-profiling to be performed,
which is an important quality control
method for ADCs that retain the
effector functions of the naked
parent antibody (ADCC). Janin-
Bussat et al. recently confirmed
that IdeS digestion of brentuximab
vedotin followed by reduction
significantly improved the peak
separation as shown by LC–MS
analysis (Figure 5[a]) (20). In
addition to the seven expected
majors peaks (Fc/2, L0, L1, Fd0, Fd1,
Fd2, and Fd3) two minor satellite
peaks were also present close to
Fd1 and Fd2 with similar masses
(Figure 5[b]). They were interpreted
as payload positional isomers by Le
et al. using the abundance of the
different DAR species (HIC data),
combined with both reversed-phase
HPLC and CE–SDS data (21).
The ultimate confirmation can be
obtained by peptide mapping based,
for example, on endoprotease Lys-C
digestion of isolated peaks and
LC–MS–MS analysis. This will be
discussed below.
Brentuximab Vedotin Positional Isomers Characterization (Bottom Level, 0.3 to 7 kDa)To confirm that peaks with the same
masses are payload positional
isomers, peaks Fd1a, Fd1b, Fd2a,
and Fd2b (Figure 5[b]) were enriched
by reversed-phase HPLC collection.
The characterization of each fraction
of interest was achieved after
endoprotease Lys-C digestion and
LC–MS analysis.
Peptide mapping of ADCs with
hydrophobic drugs linked to their
native cysteine residues by LC–MS
analysis is challenging because
the conjugated peptides are not
very soluble in aqueous buffers.
This is especially true for peptides
with two or more conjugated drugs.
As an improvement on the peptide
mapping protocol of brentuximab
vedotin (Lys-C digestion), all of the
steps including enzymatic digestion
were optimized to maintain the
hydrophobic drug-loaded peptides
in solution by the addition of solvents
(10% acetonitrile added to the
sample before digestion and 40%
isopropanol after digestion). The
data confirmed that the drug was
linked, as expected, to the inter-
chain cysteines of the heavy and
light chains. Furthermore, LC–MS–
MS confirmed the payload positional
isomers. It was unambiguously
demonstrated that the drug was
linked preferentially to the heavy
chain (HC) L15 peptide on Cys220
when only one drug was bound to
the HC. In contrast, when two drugs
were linked to the HC, they were
preferentially bound to the HC L16
peptide on Cys 226 and Cys229.
For further analysis and following
ad hoc sample preparation as
reported by Lebert et al. (22), a
similar chromatographic separation
of ADC peptides combined with
MS analysis can also be applied
to pharmacokinetic studies for
characterization of the ADCs
drug-loaded peptides distribution
after their in vivo administration.
Trastuzumab Emtansine (Top Level, Deglycosylated, 145 to 154 kDa)As demonstrated for cysteine
conjugates, HIC and native MS are
the key techniques for studying drug
distribution, the naked antibody
content, and the average DAR.
For lysine ADC conjugates on the
other hand, which are not amenable
to HIC because of their higher
heterogeneity, denaturing MS and
UV–vis spectroscopy are the most
powerful approaches.
In analogy to trastuzumab
emtansine, the ADC huN901–DM1
Advances in Biopharmaceutical Analysis – October 201528
Beck and Cianférani
IdeS
lgG(150 kDa)
F(ab)’2(100 kDa)
2 Fc/2(25 kDa)
2 Fd(26 kDa)
Fd1a71%
Fd1a26.2
Fd1b27.4
Fd2a32.5
Fd2b33.9 Fd3
39.7
Fd22.6
Fc/212.0
Fd1 Fd2
101326.0010.00 15.00 20.00 25.00 30.00 35.00 40.00 28.00 30.00 32.00
Time (min) Time (min)
PayloadCys 226
L16:Cys 229L16: Cys220
34.00
%
%
Fd1b29% LC1
19.6LC14.7
Fd2a17%
or
Fd2b83%
2 LC(25 kDa)
2 Fc/2(25 kDa)
DTT
(a)
(c)(b)
Figure 5: Subunits and positional isomers of brentuximab vedotin: (a) IdeS/Reduction workflow, (b) reversed-phase HPLC–MS profile (PLRP-S column), and (c) positional payload isomers found in Fd1 and Fd2 fractions.
Trastuzumab Emtansine-SMCC-DM1
*Trastuzumab Emtansine (SMCC)n+1-(DM1)n
* * ** *
*
Average DAR = 3.4
D328.9%
148737
D512.7%
150652
D71.8% D8
0.7%
D66.8%151610
146000 147000 148000 149000 150000 151000 152000 153000 154000
mass
0
D423.3%
149695
D17.6%146822
D01.4%
148967
D216.8%
147780
Figure 6: Deconvoluted electrospray mass spectrum of trastuzumab emtansine obtained under denaturing conditions (ESI-QTOF).
ES683362_LCESUPP1015_028.pgs 10.01.2015 21:02 ADV blackyellowmagentacyan
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ES683541_LCESUPP1015_029_FP.pgs 10.01.2015 21:53 ADV blackyellowmagentacyan
contains an average of three to four
DM1 drug molecules per huN901
IgG1 antibody molecule (same DL).
The composition of the mixture
was determined by MS of the
deglycosylated conjugate by Wang
et al. (23). Deglycosylation eliminates
the complexity in the MS spectrum
caused by the heterogeneity in the
glycosylation of the CHO cell-derived
huN901 antibody. Samples were
analyzed using size-exclusion
chromatography (SEC) coupled
on-line with electrospray ionization
(ESI) time-of-flight (TOF)-MS to
avoid salt interference with protein
ionization (24). A representative
ESI-TOF-MS spectrum of lysine
conjugated trastuzumab emtansine is
shown in Figure 6.
More recently, Marcoux et al.
reported the use of native MS
and ion mobility (IM–MS) for the
characterization of trastuzumab
emtansine, also known as T-DM1 or
Kadcyla (25). This lysine conjugate
has recently been approved for the
treatment of human epidermal growth
factor receptor 2 (HER2)-positive
breast cancer, and combines the
anti-HER2 antibody trastuzumab
(Herceptin) with the cytotoxic
microtubule-inhibiting maytansine
derivative, DM1. Native MS combined
with high-resolution measurements
and charge reduction is beneficial
for the accurate values it provides
of the average DAR and the drug
load profiles. The use of spectral
deconvolution was investigated in
detail. In addition, the use of native
IM–MS to directly determine DAR
distribution profiles and average
DAR values, as well as a molecular
modelling investigation of positional
isomers in T-DM1, was reported.
ConclusionThe development and optimization
of ADCs is increasingly being driven
by a need to improve its analytical
and bioanalytical characterization
by assessing the main ADC quality
attributes: drug load distribution,
amount of naked antibody, and
average DAR. These needs have
been recently fulfilled by a number
of cutting-edge MS methods and
optimized workflows used at different
levels.
At the top level, native MS and
native IM–MS is successfully used
in addition to HIC, the reference
method for quality control of
inter-chain cysteinyl-linked ADCs.
At the middle level, reduced or
IdeS-digested ADCs are analyzed
by LC–MS, which is used as an
orthogonal method to gain structural
insights on ADC subunits. In
addition, the use of IdeS allows the
analysis of the Fc/2 that has been
separated from the Fd fragment and
the light chain. As a result, the full
glyco-profiling and demonstration
of the absence of additional
conjugation are easily achieved.
At the bottom level, improved ADC
LC–MS peptide mapping methods
used to characterize the drug-loaded
peptides and to identify positional
isomers at cysteine residues have
been developed. All the steps of
the method including enzymatic
digestion have been optimized
to maintain the hydrophobic
drug-loaded peptides in solution by
the addition of solvents.
AcknowledgementsThe authors acknowledge F. Debaene
and J. Marcoux (LSMBO, Strasbourg,
France) and E. Wagner-Rousset, M.C.
Janin-Bussat, O. Colas, M. Excoffier, L.
Morel-Chevillet, C. Klinguer-Hamour,
and T. Champion (CIPF, St-Julien en
Genevois, France) for their contribution
to the development of new ADC
characterization methods.
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Hamour, M. Jaquinod, and A. Beck,
Bioanalysis 7, 1237–1251 (2015).
(23) A.C. Lazar, L. Wang, W.A. Blattler, G.
Amphlett, J.M. Lambert, and W. Zhang,
Rapid Commun. Mass Spectrom. 19,
1806–1814 (2005).
(24) L. Wang, G. Amphlett, W.A. Blattler, J.M.
Lambert, and W. Zhang, Protein Sci. 14,
2436–2446 (2005).
(25) J. Marcoux, T. Champion, O. Colas, E.
Wagner-Rousset, N. Corvaia, A. Van
Dorsselaer, A. Beck, and S. Cianferani,
Protein Sci. 24, 1210–1223 (2015).
Dr. Alain Beck is Senior Director,
Antibody/ADC Physico-Chemistry
and member of the board of directors
of the Centre d’Immunologie Pierre-
Fabre. He has contributed to the R&D
of anticancer mAbs (in collaboration
with Merck and Abbvie), vaccines,
and peptides. He is associate editor
of mAbs, an inventor of 16 patents,
author of more than 140 publications
and reports, and he has contributed
to more than 200 meetings.
Dr. Sarah Cianférani is CNRS
Research Director and Director of
the BioOrganic Mass Spectrometry
Laboratory (LSMBO) at the IPHC,
University of Strasbourg, France. Her
research is focused on developments
and applications of advanced native
MS, IM–MS, and HDX-MS methods
for biological non-covalent complex
characterization.
Advances in Biopharmaceutical Analysis – October 201530
Beck and Cianférani
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31www.chromatographyonline.com 31www.chromatographyonline.com
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Since its invention in the early 20th
century (1), mass spectrometry (MS)
has been used to discover new
chemical elements and their isotopes
(2), explore martian soil for organic
matter (3), and study biological
processes by profiling proteomes or
metabolomes (4,5). Biopolymers, and
especially proteins, are the subject
of intensive investigation. They are
characterized by different levels of
structural organization, ranging from
the primary structure represented
by the amino acid sequence, over
secondary structural elements such
as alpha-helices and beta-sheets,
and the three-dimensional orientation
of the polypeptide chain, and finally
to the assembly of subunits into
protein complexes.
Analysis of protein structure
using MS was first possible in the
mid 1980s, when the soft ionization
techniques of electrospray ionization
(ESI) (6) and matrix-assisted laser
desorption–ionization (MALDI) (7)
were introduced. Implementation
of MS to protein analysis initially
focused on large-scale identification
(8) and the determination or
confirmation of primary structure
(9), whereas newer technologies
have laid the ground for the study
of tertiary and even higher order
structures of protein molecules and
complexes.
The analysis of biopharmaceuticals
(therapeutic proteins developed
for disease treatment) requires
analytical techniques able to
elucidate the various structural levels
to ensure their efficacy and safety
in patients. Several MS techniques
are indispensable in the toolbox of
physico-chemical characterization
methods available for the analysis
of therapeutic proteins (10). Some of
the methods used in the elucidation
of higher order structural elements
of proteins — including native MS,
ion mobility MS, hydrogen-deuterium
exchange MS, and chemical
cross-lining MS — will be discussed
in this feature article.
Native Mass Spectrometry Experiments using electrospray
ionization mass spectrometry
(ESI–MS) for the analysis of intact
proteins were first performed by
J.B. Fenn’s group (6). The study
used strongly denaturing conditions
created by using 50–90% organic
solvent containing acetic acid
or trifluoroacetic acid and was
beneficial for the detection of
multiple-charge protein ions with a
quadrupole mass spectrometer of
m/z 1500 upper nominal mass limit.
However, it was soon discovered
that the charge-state distribution
of electrosprayed proteins was
significantly influenced by the
protein structure prevalent under
non-denaturing or denaturing
conditions (11). This then led to the
implementation of native MS.
Native MS aims to maintain
the three-dimensional structure
of proteins or protein complexes
as much as possible during an
experiment (12) by using conditions
that reflect the protein’s native
environment. The overall charge
of a protein ion is limited by the
number of ionizable functionalities
that are accessible on the surface
for charging, predominantly through
protonation or deprotonation, leading
to the observation of low-charged
species in mass spectra. Weakly
bound, noncovalent complexes
— including proteins interacting
with inhibitors, cofactors, metal
ions, carbohydrates, or peptides
— can be preserved during the
electrospray process facilitating the
study of the structure, stoichiometry,
and association constant of such
biomolecular complexes (13). The
reduction of charge requires the
use of mass spectrometers with
an extended mass range such as
time-of-flight (TOF) (14), or more
recently orbitrap (15) mass analyzers
to detect the low-charged protein
species.
Trastuzumab (INN; trade name
Herceptin) is a monoclonal antibody
that interferes with the human
epidermal growth factor receptor
2 (HER2) and is used to treat
HER2-positive breast cancers.
Figure 1(a) and 1(c) illustrate the
difference between mass spectra
of trastuzumab when analyzed
under (a) denaturing versus (c)
Higher Order Mass Spectrometry Techniques Applied to BiopharmaceuticalsChristian G. Huber, Department of Molecular Biology, Division of Chemistry and Bioanalytics, and Christian Doppler
Laboratory for Innovative Tools for Biosimilar Characterization, University of Salzburg, Salzburg, Austria.
The recent trends in mass spectrometric techniques — including native mass spectrometry (MS), ion mobility spectrometry (IMS), hydrogen–deuterium exchange MS (HDX MS), and chemical cross-linking MS (CXMS) — employed to elucidate higher-order structures of protein complexes and the practical implementations of these methods are discussed.
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32 Advances in Biopharmaceutical Analysis – October 2015
Huber
non-denaturing conditions. Under
denaturing conditions (a) charge
states from 33+ to 60+ are detected
in an m/z range of 2200–4500,
whereas non-denaturing conditions
(c) yield charge states of 22+ to 28+
at m/z 5200–6700. Deconvolution of
both mass spectra gives equivalent
masses for the uncharged species
with masses in the range of 147–148
kDa and also reveals several different
protein species that represent
the different glycoforms of the
monoclonal antibody. Such analysis
can therefore readily reveal the
glycosylation pattern of the protein, a
highly important quality parameter of
recombinant biopharmaceuticals.
Native MS has been shown to be
a highly efficient tool for determining
binding stoichiometry of a
monoclonal antibody with its antigen.
Humanized murine monoclonal
antibody (hzmAb), directed against
the junctional adhesion molecule
A (JAM-A) to have antiproliferative
and antitumoural properties,
was titrated with its antigen and
then analyzed using native MS
to reveal non-covalent complex
stoichiometries (17). Three species
were detected when equimolar
amounts of antibody and its
target antigen were incubated
including free antibody, 1:1, and 1:2
antibody:antigen complex. Two molar
equivalents of antigen led to an
almost quantitative formation of the
1:2 complex, while eightfold molar
excess yielded a 1:4 complex with a
small portion of 1:3 complex.
Ion Mobility SpectrometryDeveloped in the 1960s, ion
mobility spectrometry (IMS)
enables the generation of size- and
conformation-dependent information
that is not possible using MS alone.
When coupled to mass spectrometry
this technique has the potential
bc
a
1000
2000
4000
6000
8000
4000
2000 3000 4000
3000 4000 5000 6000 7000 8000
m/z
m/z
5 10
Drift Time (millisec)
15 20 25
5 10
Drift Time (millisec)
15 20 25 30
d
e
(a) (b)
(c) (d)
45+
42+
23+27+
25+
Figure 1: MS and IMS analysis of intact trastuzumab. (a) and (c): Intact MS analysis of trastuzumab. ESI-TOF mass spectra of trastuzumab in denaturing (a) or native (c) conditions. The inserts shows an extended view of the 44+ (a) and 25+ (c) charge states with resolution of the different glycoforms: (a) 147 917.1 ± 1.1 Da (G0/G0F), (b) 148 061.7 ± 0.8 Da (G0F/G0F), (c) 148 222.4 ± 0.9 Da (G0F/G1F), (d) 148 383.8 ± 0.8 Da (G1F/G1F), and (e) 148 544.3 ± 1.0 Da (G1F/G2F). (b) and (d): IMS analysis of trastuzumab. Ion mobility mobilograms of trastuzumab in denaturing (b) or native (d) conditions. IMS data obtained in native conditions (d) reveal small amounts of dimeric mAb. Adapted and reproduced with permission from Analytical Chemistry 84, Alain Beck, Sarah Sanglier-Cianférani, and Alain Van Dorsselaer, Biosimilar, biobetter, and Next Generation Antibody Characterization
by Mass Spectrometry, 4637–4646 (2012) © American Chemical Society.
ESI -sourceIon
guide Quadrupole TWIMS cell
Trap Transfer
Pusher Detector
Refectron
Figure 2: Schematic diagram of a quadrupole-traveling wave ion mobility–time of flight instrument.
ES683381_LCESUPP1015_032.pgs 10.01.2015 21:03 ADV blackyellowmagentacyan
33www.chromatographyonline.com
Huber
to separate isomers, isobars, and
conformers; significantly reduce
chemical background; and detect
aggregates of biopharmaceuticals.
IMS separation of ions is possible
using differing separation powers,
analyte detection, and hyphenation
to mass spectrometry (18)
including: (1) drift-time ion mobility
spectrometry (DTIMS); (2) aspiration
ion mobility spectrometry (AIMS);
(3) field-asymmetric waveform ion
mobility spectrometry (FAIMS);
or (4) travelling-wave ion mobility
spectrometry (TWIMS). DTIMS
and TWIMS are the two principles
most often used in commercial
instruments. In a DTIMS device ions
are moved through a uniform, linear-
field drift tube filled with a so-called
“buffer gas” through a small, uniform
electric field. The moving ions are
attenuated by collisions with the
buffer gas depending on their overall
charge and collision cross-section
(18). Ions with multiple charges and
a small cross-sectional area move
faster through the drift cell than
low-charge ions with large collisional
cross-sections. TWIMS is a type of
IMS that utilizes travelling waves
created by a series of ring-shaped
electrodes that split the structure of
the drift cell into a series of segments
(Figure 2). Instead of a uniform
linear field, a high field is applied
to one segment of the cell that is
subsequently swept through the
cell in the direction of ion migration.
Consequently, movement and
separation of ions in the mobility cell
is accomplished by means of pulses
of an electric field passing through.
An example for the application of
TWIMS to the analysis of monoclonal
antibodies is illustrated in Figure 1(b)
and (d). IMS analysis of trastuzumab
under denaturing conditions reveals
a large number of highly charged
species clustering at drift times
between 10 and 15 ms, while native
conditions clearly distinguish
between the different, low-charged
species in a drift time range of
7–25 ms. This contrast in drift
behaviour is advantageous for the
analysis of more complex mixtures
of biopharmaceuticals, particularly
when looking at sequence variants or
other post-translational modifications
such as oxidation or pyroglutamate
formation.
Another practical example
of IMS characterization of
biopharmaceuticals (less directed
towards higher order elucidation) is
outlined in Figure 3. Here, a reduced
mouse monoclonal antibody (IgG1,
κ) sample comprising of both heavy
and light chains was introduced
into an ESI-quadrupole-IMS-TOF
system (19). The two-dimensional ion
mobilogram-mass spectrum depicted
in Figure 3(a) clearly shows that light
and heavy chains can be readily
separated as different species
without any other upfront separation
technique. Multiple charged species
related to the light and heavy
chains were differentiated between
using mass spectra extracted from
the encircled areas in Figure 3(a)
without mutually interfering signals.
Extracted mass spectra (Figure 3[b]
and [d]) were deconvoluted using a
maximum entropy algorithm, yielding
spectra of uncharged species, as
illustrated in Figure 3(c) and (e).
As expected, the light chain is
detected as a single species, while
the spectrum of the heavy chain
reveals at least three glycoforms,
characterized by different galactose
content in the attached N-glycan
(162 Da mass difference). This
example nicely demonstrates the
benefits of an additional dimension
of separation, although an upfront
Light Chain
Heavy Chain
2000
1500
1000
700
10026+
(b)
(a)
(d)
(e)(c)
28+ 23+
20+
17+
52+
45+
41+
36+
800 1200 1600 2000 2400 28000
%
100
024050 24150 24250 24350 24450 49600 49800 50000 50200 50500
massmass
50249
49779
49924
50086
** *
*
*
*** *
2422624177
24199
%
100
0
%
100
800 1200 1600 2000 2400 2800
m/zm/z
0
%
3.2 6.4
Drift Time (millisec)
m/z
9.6 12.8
Figure 3: On-line LC–MS analysis of a completely reduced IgG1 antibody using ion mobility-TOF mode. (a) Two-dimensional plot of ion drift time vs. m/z for the reduced IgG1 obtained using the ion mobility separation (7.5V pulse). (b) Combined raw mass spectrum of the light chain. (c) Deconvoluted mass spectrum of the light chain. (d) Combined raw mass spectrum of the heavy chain. (e) Deconvoluted mass spectrum of the heavy chain. Adapted and reproduced with permission from Rapid Communications in Mass Spectrometry 22, Petra Olivova, Weibin Chen, Asish B. Chakraborty, and John C. Gebler, Determination of N-glycosylation sites and site heterogeneity in a monoclonal antibody by electrospray quadrupole ion-mobility time-of-flight mass spectrometry, 29–40 (2007) © John Wiley and Sons.
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34 Advances in Biopharmaceutical Analysis – October 2015
Huber
separation method such as high
performance liquid chromatography
(HPLC) or capillary electrophoresis
(CE) may be necessary — especially
for the quantitative analysis of trace
amounts of impurities that is often
indispensable for quality control.
Hydrogen-Deuterium Exchange Mass SpectrometryThe native state of proteins is
generally characterized by a tightly
folded, compact structure that
exposes a well-defined surface
to its environment. Denaturation
under conditions — such as high
temperature, extreme pH, adsorption
to surfaces, or dissolution in organic
solvent — results in the protein
unfolding and forming a significantly
different surface. Similarly,
interactions of a protein with other
molecules such as small drugs,
nucleic acids, or other proteins
can lead to a significant change in
surface properties.
Denaturing and complex formation
can also have a profound influence
on the exchangeability of protons at
the surface of a protein molecule —
the acidic protons of the carboxyl
groups or acidic side chains
of aspartate and glutamate are
normally the most easily and rapidly
exchanged while the amide protons
of the protein backbone are much
less prone to exchange. Exchange
can be monitored by dissolution
of a protein in heavy water (D2O),
which leads to a hydrogen exchange
by deuterium (H/D exchange)
in a few minutes to hours. In
proteins, the exchange rates for the
different hydrogen atoms strongly
depend on accessibility and
therefore on protein conformation
or association into higher order
structures. The substitution of
exchangeable hydrogen atoms with
deuterium atoms forms the basis of
hydrogen-deuterium exchange mass
spectrometry (HDX-MS) (20).
A schematic workflow of HDX-MS
is depicted in Figure 4. In brief, a
protein with exchangeable hydrogens
is dissolved for different periods
of time at ambient or elevated
incubation temperature (20–40
°C) in deuterated water (buffered
to pH around 7). Depending on
exchangeability, hydrogen atoms
are replaced by deuterium atoms
during the incubation time, before
the exchange is quenched upon
acidification and cooling to 0
°C. Proteins are then digested
under quenching conditions,
and the resulting peptides are
separated by low-temperature
HPLC, and finally analyzed by
tandem mass spectrometry (MS–
MS) upon fragmentation either by
collision-induced dissociation (CID)
or electron-transfer dissociation
(ETD). Characteristic mass shifts
in the fragment ions are indicative
for the presence and position(s) of
deuterium atoms. Analysis of the
kinetics of deuterium uptake yields
information about the accessibility
of hydrogens at different positions
in the protein, which allows valuable
insights into the three-dimensional
structure of proteins or protein
complexes.
HDX-MS has been successfully
used to compare three-dimensional
structures of biopharmaceuticals,
which is essential to demonstrate
manufacturing consistency
to regulatory agencies or
provide a proof of structural
equivalence between an originator
biopharmaceutical and its biosimilar.
The advantage of this approach is
that it probes the whole molecule
instead of just certain substructures.
The results of an interrogation of
the three-dimensional structure of
interferon-β-1a, a 20-kDa cytokine
used to treat multiple sclerosis
(traded under the names Avonex
(Biogen), Rebif (Merck Serono or
Pfizer), or CinnoVex (CinnaGen) as
a biosimilar), are shown in Figure
5 (22). Hydrogen exchange rates
determined for five different peptic
peptides effectively show the impact
of different manufacturing conditions
as well as post-translational
modifications — modification with
poly(ethylene glycol) (PEG); or
oxidation at C17, M1, M36, M62, and
M117 — on protein structure.
No significant alteration in
hydrogen-deuterium exchange
profile was observed, even though
production involved different
batches, using different cell media,
and was subjected to N-terminal
modification with PEG. A significant
impact was however found for
methionine or cysteine oxidation,
and because the oxidized peptides
incorporated more deuterium
compared to the reference
analogues, it could be concluded
that oxidized interferon-β-1a is more
solvent exposed and less hydrogen
bonded.
Although this example convincingly
demonstrated the applicability of
H HH H
Z7
t1
H/D exchange
Quench
(pH 2.5, 0oC )
(pepsin, pH 2.5, 0oC )
Cooled
LC-MS
Time
co
nte
nt
Time
(protein)
Gas-phase
Solution-phase
cleavage
cleavage
(peptide)
Gas-phase
cleavage
D2O
t2t3t4
C7
C6
C5
C4
C3
C2
C1
R1
N
O
NN
NN
NN
NOH
OOOO
O O O
2
R3
R5
R7
R8
R6
R4
R2
Z6
Z5
Z4
Z3
Z2
Z1
H
H
H
H
H
HH
H
HH
H
H
H
H
HH
H
H
H
H
HH
H H
HH
H H
H
H
HH H
H
H
D
DD
D
DD
D
D
DDD
D
D
D D
D
D
H
HH
H
H
H
H H
H
H
HH H
H
H
D
DD
D
DD
D
D
DDD
D
D
co
nte
nt
D
D
D
H
HH
H
H
H
HH
H
H
HH H
H
H
D
DD D
D D
D
D
DDD
D
D
D
H
HH
H
H
H
Figure 4: Principle of hydrogen/deuterium exchange mass spectrometry. Adapted and reproduced with permission from Accounts of Chemical Research 47, Kaspar D. Rand, Martin Zehl, and Thomas J.D. Jørgensen, Measuring the Hydrogen/Deuterium Exchange of Proteins at High Spatial Resolution by Mass Spectrometry: Overcoming Gas-Phase Hydrogen/Deuterium Scrambling, 3018–3027 (2014) © American Chemical Society.
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35www.chromatographyonline.com
Huber
HDX-MS for revealing structural differences in homogenous biopharmaceuticals, the authors also pointed out that it is not capable, at the moment, to detect conformational differences in coexisting, low-level (<10%) components of the population (22).
Chemical Cross-Linking Mass Spectrometry (CXMS)Three-dimensional structures of proteins can be determined with atomic resolution by using high-resolution methods such as X-ray crystallography or nuclear magnetic resonance (NMR)
spectroscopy, but the high amount of sample required for these methods (typically in the milligram range) makes them impractical for biological studies. In comparison, low-resolution structural data generated by chemical cross-linking MS (CXMS) uses much less sample amounts (in the nanogram to picogram range) to generate highly valuable data (23). Low-resolution structural information is obtained by chemically cross-linking functional groups in a protein by means of a bifunctional cross-linker, which gives information about the distance of the cross-linked functional groups
in a protein molecule or a protein complex.
The most common functional groups available for cross-linking in proteins are the lysine amino groups. Sulphydryl groups of cysteines are another possibility, but they can become involved in the three-dimensional structure of a protein particularly when created by reduction of disulphide-bridges in the native protein. Although formaldehyde is the oldest cross-linking reagent, the most commonly utilized reagents are based on bifunctional N-hydroxy-succinimide esters, which readily react with free amino groups (and in a side reaction also with hydroxyl groups of tyrosine) to create a stable amide or imide bond upon release of N-hydroxysuccinimide. Depending on the length of the cross-linking spacer, different distances of amino acids can be probed, ranging from (almost) zero for formaldehyde to 6.4 Å for disulphosuccinimidyl tartrate, 11.4 Å for bis(sulphosuccinimidyl)suberate (10 atoms), and 16.1 Å for ethylene glycol bis(sulphosuccinimidyl succinate (14 atoms) (24).
Figure 6 shows an outline of a cross-linking experiment. After the formation of intramolecular or intermolecular cross-links, the protein or protein complex is proteolytically digested and the resulting peptides are analyzed via HPLC–MS–MS. Because the crosslink remains unaffected by the proteolysis, cross-linked amino acids are revealed through the corresponding cross-linked peptides. To more easily identify cross-linked products, isotope-labelled cross-linking reagents with 50% heavy isotope exchange can be used. Thus, cross-linked peptides are recognized by 1:1 doublets of mass signals for the light and heavy versions, which is usually achieved with the help of computer-based searching algorithms (25). The distance information obtained from the cross-linking experiment is then used to build and verify structural models for proteins or protein complexes.
Chemical cross-linking can also be utilized to directly analyze stabilized protein complexes. For
1. (8 - 15)FLQRSSNF
7
6
5
4
3
2
1
00 1 10 100 1000
5
4
3
2
1
00 1 10 100 1000
12
10
8
6
4
2
00 1 10 100 1000
12
10
8
6
4
2
00 1 10 100 1000
5678
43210
0 1 10 100 1000
7
6
5
4
3
2
1
00 1 10 100 1000
5
4
3
2
1
00 1 10 100 1000
12
10
8
6
4
2
00 1 10 100 1000
12
10
8
6
4
2
00 1 10 100 1000
5678
43210
0 1 10 100 1000
7
6
5
4
3
2
1
00 1 10 100 1000
5
4
3
2
1
00 1 10 100 1000
12
10
8
6
4
2
00 1 10 100 1000
12
10
8
6
4
2
00 1 10 100 1000
5678
43210
0 1 10 100 1000
7
6
5
4
3
2
1
00 1 10 100 1000
5
4
3
2
1
00 1 10 100 1000
12
10
8
6
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2
00 1 10 100 1000
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Rela
tive a
bu
nd
an
ce (
Da)
Rela
tive a
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nd
an
ce (
Da)
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tive a
bu
nd
an
ce (
Da)
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tive a
bu
nd
an
ce (
Da)
2. (62 - 67)MLQNIF
3. (88 - 101)LANVYHQINHLKTV
4. (121 - 134)HLKRYYGRILHYLK
5. (154 - 162)FYFINRLTG
(a)Differentbatches
(c)Different
media
(b)Pegylated
(d)Oxidized
1. (8- 15)
Time (min) Time (min) Time (min) Time (min) Time (min)
3. (88- 101)
5. (154- 162)2. (62- 67)
4. (121- 134)
Figure 5: Deuterium incorporation graphs generated for five interferon-β-1a (IFN) peptic peptides from four different H/DX MS comparability experiments. In each graph, the reference IFN data are the black lines with closed triangles, while the experimental IFN data, to which it is being compared, is the red line with open circles. Row a: Comparison of two different large scale IFN batches prepared over eight years apart; row b: Comparison of IFN versus N-terminally PEGylated IFN; row c: Comparison of IFN produced using different cell culture media and growth conditions; row d: Comparison of IFN versus oxidized IFN (oxidation of Met and Cys residues C17, M1, M36, M62, and M117 was 100%). Adapted and reproduced with permission from Journal of Pharmaceutical Science 100, Damian Houde, Steven A. Berkowitz, and John R. Engen, The utility of hydrogen/deuterium exchange mass spectrometry in biopharmaceutical comparability studies, 2071–2086 (2010) © John Wiley and Sons.
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36 Advances in Biopharmaceutical Analysis – October 2015
Huber
example, disuccinimidyl suberate,
as well as 1,1’-(suberoyldioxyl)
bisazabenzotriazole) were used as
cross-linkers to stabilize the complex
between the bovine prion protein
and a specific antibody against
it, the antibody 3E7 (26). Direct
analysis of the reaction products by
matrix-assisted laser desorption–
ionization MS revealed both the
free prion protein and the free
antibody together with 1:1 and 1:2
antibody:prion protein complexes.
ConclusionsIn conclusion, MS, traditionally
regarded as one of the most
important analytical methods for
the determination of the primary
structure of proteins, is increasingly
contributing to the elucidation of
diverse fundamental aspects of
the tertiary and even quaternary
structure of proteins and protein
complexes. In spite of providing
less spatial resolution, the major
strength of MS-based investigations
over NMR spectroscopy or X-ray
crystallography lies within the
comparatively low amounts of
sample required for successful
analysis, typically a few picograms
to nanograms. Such studies
are, however, only feasible with
substantial support through elaborate
computational algorithms and
workflows, which requires significant
involvement of bioinformatics into
data evaluation.
AcknowledgementsThe financial support by the Austrian
Federal Ministry of Economy, Family,
and Youth, the National Foundation
of Research, Technology, and
Development, and by a Start-up Grant
of the State of Salzburg is gratefully
acknowledged.
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60, 2299–2301 (1988).
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Yates, J. Am. Soc. Mass Spectrom. 5,
976–989 (1994).
(9) H. Nau and K. Biemann, Abstracts of
Papers of the American Chemical Society
62–62 (1974).
(10) R.J. Falconer, D. Jackson-Matthews,
and S.M. Mahler, J. Chem. Technol.
Biotechnol. 86, 915–922 (2011).
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Biomed. Environ. Mass Spectrom. 17,
411–414 (1988).
(12) M. Przybylski and M.O. Glocker, Angew.
Chem. Int. Ed. 35, 806–826 (1996).
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175–186 (2000).
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Standing, C. P. Whitman, and S. B. Kent,
Proc. Natl. Acad. Sci. U.S.A. 93, 6851–
6856 (1996).
(15) S. Rosati, R.J. Rose, N.J. Thompson,
E. van Duijn, E. Damoc, E. Denisov, A.
Makarov, and A. J. R. Heck, Angew.
Chem.-Int. Ed. 51, 12992–12996 (2012).
(16) A. Beck, S. Sanglier-Cianferani, and A.
Van Dorsselaer, Anal. Chem. 84, 4637–
4646 (2012).
(17) C. Atmanene, E. Wagner-Rousset, M.
Malissard, B. Chol, A. Robert, N. Corvaia,
A. Van Dorsselaer, A. Beck, and S.
Sanglier-Cianferani, Anal. Chem. 81,
6364–6373 (2009).
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H.H. Hill, J. Mass Spectrom. 43, 1–22 (2008).
(19) P. Olivova, W. Chen, A.B. Chakraborty,
and J.C. Gebler, Rapid Commun. Mass
Spectrom. 22, 29–40 (2008).
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Soc. 115, 6317–6321 (1993).
(21) K.D. Rand, M. Zehl, and T.J.D. Jorgensen,
Acc. Chem. Res. 47, 3018–3027 (2014).
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J. Pharm. Sci. 100, 2071–2086 (2011).
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(2003).
(25) A. Leitner, T. Walzthoeni, and R. Aebersold,
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Anal. Chem. 82, 172–179 (2010).
Christian Huber was educated as an
analytical chemist from 1985 to 1993
at the University of Innsbruck, Austria.
Following a lecturing qualification at
the University of Innsbruck in 1997, he
held the chair of analytical chemistry
position at Saarland University in
Germany from 2002 to 2008. In
2008, he was made a professor of
chemistry for biosciences at the
Department of Molecular Biology of
the University of Salzburg, Austria.
His research interests include
bioanalytical chemistry and proteome
and metabolome analysis, as well
as in-depth characterization of
therapeutic proteins.
Proteolysis
LC-MS/MS
Cross-linking
Map of cross-links
Intra and inter-chainSelection of structural models cross-links
Set of distance restraints
Protein complexProtein 1
Protein 2
Protein 3
Figure 6: Schematic of a workflow for chemical cross-linking of protein complexes followed by digestion and analysis by HPLC–MS–MS. Reproduced with permission from http://daltonlab.iqm.unicamp.br/research.html.
ES683396_LCESUPP1015_036.pgs 10.01.2015 21:03 ADV blackyellowmagentacyan
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Ph
oto
Cre
dit: G
IPh
oto
Sto
ck/G
ett
y Im
ag
es
The development of protein‑based
pharmaceuticals, or biopharmaceuticals,
is by far the fastest growing part of
the pharmaceutical industry today.
With over 1500 biopharmaceuticals
in clinical development and more and
more companies shifting their R&D
efforts towards this sophisticated and
relatively profitable class of drugs,
the pharmaceutical landscape has
changed beyond recognition compared
to 20 or even 10 years ago. As a result,
the field of bioanalysis that supports
drug development by measuring the
concentrations of drugs or relevant
endogenous molecules in biological
samples has also seen many changes.
The quantitative determination of
biopharmaceuticals has traditionally
been the domain of ligand‑binding
assays, such as ELISA. However, in the
past few years there has been a clear
increase in the application of alternative
analytical platforms, in particular liquid
chromatography coupled to tandem
mass spectrometry (LC–MS–MS),
which has been the workhorse for
small‑molecule bioanalysis for over 20
years (1–5).
Over the past decade, there
have been many advances in the
LC–MS–MS‑based quantitation of
biopharmaceuticals, both from an
analytical and a conceptual point
of view. In this article, an overview
is given of the many aspects of
this field of analytical research by
reference to a selection of recent
applications.
Protein DigestionTandem mass spectrometry remains
the detection technique of choice
for the quantitative determination
of biopharmaceuticals because of
its sensitivity and its widespread
availability in the pharmaceutical
and related industries. However,
the use of LC–MS–MS to quantify
biopharmaceuticals is more complex
than for small molecules because
it is not directly compatible with
molecules with a mass above
around 5000 Da. The ions of larger
analytes are distributed over many
different charge states and usually
do not readily fragment and this
considerably reduces sensitivity.
Therefore, a typical step in the
analysis is the (enzymatic) digestion
of a biopharmaceutical into a
mixture of smaller peptides, followed
by the analysis of the digest and
quantitation of one or more so‑called
signature peptides as a measure for
the intact protein. Digestion is usually
performed using the enzyme trypsin,
which cleaves the amino acids chain
in proteins after a lysine or arginine.
Trypsin is popular because it is
readily available at a reasonable
price and can cleave proteins
into peptides of a size (500–2000
Da) that is well suited for MS–MS
detection.
Protein digestion enormously
increases the complexity of a
biological sample. Matrices such as
plasma contain proteins at a total
concentration of around 80 mg/mL
and, when no further clean‑up of
the sample is performed, each of
these proteins is cleaved into a
series of peptides that are all of a
similar size and have more or less
comparable physicochemical and
analytical properties. Therefore, it
is often challenging to detect low
concentrations of a signature peptide
in a digest, because of the presence
of so many endogenous peptides,
which all consist of combinations of
the same 20 amino acids and often
occur at much higher levels than the
signature peptide itself.
Despite the selective nature of
MS–MS detection, chromatograms
of digested biological samples
often contain many background
Advances in Liquid Chromatography–Tandem Mass Spectrometry (LC–MS–MS)‑Based Quantitation of Biopharmaceuticals in Biological SamplesNico C. van de Merbel, Bioanalytical Laboratory, PRA Health Sciences, Assen, The Netherlands and Analytical Biochemistry,
Department of Pharmacy, University of Groningen, The Netherlands.
Liquid chromatography coupled to tandem mass spectrometry (LC–MS–MS) has recently become a more and more popular alternative to traditional ligand-binding assays for the quantitative determination of biopharmaceuticals. LC–MS–MS offers several advantages such as improved accuracy and precision, better selectivity, and generic applicability without the need for raising analyte-directed antibodies. Here we discuss the technical requirements for a successful LC–MS–MS method for the quantitation of biopharmaceuticals and evaluate the advantages and disadvantages compared to ligand-binding assays.
38 Advances in Biopharmaceutical Analysis – October 2015
ES683326_LCESUPP1015_038.pgs 10.01.2015 20:52 ADV blackyellowmagentacyan
peaks originating from endogenous
peptides that show a response at
the mass transition of the signature
peptide. Figure 1 shows this effect
for a fixed concentration of digested
salmon calcitonin in the presence
of increasing amounts of digested
plasma (6). The selectivity of the
method is clearly affected by the
presence of endogenous background
peptides. As a result, method
sensitivity is also heavily impacted
— in this case the achievable lower
limit of quantitation (LLOQ) increases
100‑fold, from 0.2 ng/mL (60 pM) in
the absence of matrix peptides to
20 ng/mL (6 nM) in the presence of
50% of digested plasma.
A review of current literature (1,4)
shows that a typical LLOQ for a
biopharmaceutical in plasma or
serum, only treated by digestion,
is in the high ng/mL to low µg/mL
range (corresponding to low nM
levels for many proteins). Figure 2
shows an example chromatogram for
a signature peptide of recombinant
human alpha‑glucosidase at its
LLOQ of 0.5 µg/mL (5 nM) in human
plasma (7).
39www.chromatographyonline.com
van de Merbel
100
0
1.75 2.00 2.25 2.50
2.94
0%
1%
5%
10%
20%
50%
2.93
1.84 2.644.24
3.723.583.22
2.922.64
2.50
2.332.162.041.76
2.162.33
2.50
2.54
2.91
3.23 3.74 4.23 4.54 4.86 5.055.44
5.40
5.754.944.514.384.20
3.723.54
3.21
2.902.81
2.47
2.302.14
2.031.76
2.00
2.22
2.74
3.043.19
3.453.66 4.20 4.41
4.72
5.24
1.76 1.932.04
5.454.874.554.24
2.75 3.00 3.25 3.50 3.75 4.00 4.25 4.50 4.75 5.00 5.25 5.50 5.75 6.0
1.75 2.00 2.25 2.50 2.75 3.00 3.25 3.50 3.75 4.00 4.25 4.50 4.75 5.00 5.25 5.50 5.75 6.0
1.75 2.00 2.25 2.50 2.75 3.00 3.25 3.50 3.75 4.00 4.25 4.50 4.75 5.00 5.25 5.50 5.75 6.0
1.75 2.00 2.25 2.50 2.75 3.00 3.25 3.50 3.75 4.00 4.25 4.50 4.75 5.00 5.25 5.50 5.75 6.0
1.75 2.00 2.25 2.50 2.75 3.00 3.25 3.50 3.75 4.00 4.25 4.50 4.75 5.00 5.25 5.50 5.75 6.0
1.75 2.00 2.25 2.50 2.75 3.00 3.25 3.50
Time (min)
Time (min)
Time (min)
Time (min)
Time (min)
Time (min)
3.75 4.00 4.25 4.50 4.75 5.00 5.25 5.50 5.75 6.0
%
100
0
%
100
0
%
100
0
%
100
0
%
100
0
%
Figure 1: LC–MS–MS (m/z 561.9 to m/z 204.0) chromatograms of a signature peptide of 2 ng/mL salmon calcitonin in samples containing increasing amounts of human plasma digest. Analyte peak at 2.9 min. Adapted and reproduced with permission from Analytical Chemistry 85, K.J. Bronsema, R. Bischoff, and N.C. van de Merbel, High-Sensitivity LC–MS/MS Quantification of Peptides and Proteins in Complex Biological Samples: The Impact of Enzymatic Digestion and Internal Standard Selection on Method Performance, 9528–9535 (2013) © American Chemical Society.
ES683321_LCESUPP1015_039.pgs 10.01.2015 20:53 ADV blackyellowmagentacyan
Signature Peptide SelectionThe possibilities for selecting
a proper signature peptide are
usually rather limited. First and
foremost, it is essential that the
selected signature peptide has a
unique amino acid sequence that
does not naturally occur in any of
the endogenous matrix proteins.
Selection of a non‑unique signature
peptide results in an overestimation
of analyte concentrations, because
the same amino acid sequence
that is released from endogenous
proteins would contribute to the
overall signal. This often disqualifies
a large number of the theoretical
signature peptides, particularly for
biopharmaceuticals with a high
degree of similarity to endogenous
proteins, such as humanized
antibodies (if these need to be
quantified in human plasma). In
addition, other criteria are applied to
ensure robustness of the LC–MS–MS
assay. Peptides containing unstable
amino acids, such as methionine
and tryptophan that can be oxidized,
or glutamine and asparagine that
can be deamidated, are usually
disregarded to avoid losses
during analysis, although forced
oxidation of a signature peptide to
a stable oxidized product has been
successfully used (8). Similarly,
peptides with (variable) post‑
translational modifications — such
as O‑ or N‑glycosylated amino acids
— are typically excluded because
these would introduce undesirable
heterogeneity. In addition, peptides
that are too small, too large,
too polar, or too hydrophobic
might cause analytical problems
because of adsorption, sub‑optimal
chromatographic behaviour, or
limited selectivity and sensitivity.
In the end, there may be just a few
out of the many potential signature
peptides that can be successfully
used in practice.
Protein ExtractionAn obvious way to improve selectivity
and sensitivity of an LC–MS–MS
method is to remove interfering
matrix proteins prior to digestion,
which can be achieved by applying
immunocapture (IC) techniques.
Magnetic beads or other resins are
coated with a protein that displays
a high binding affinity towards
the analyte, typically an antibody
raised against the analyte or the
pharmacological target to which a
biopharmaceutical is directed. By
mixing the sample with a suspension
of the beads or passing it through
a cartridge filled with the resin, the
analyte is selectively isolated from
the complex sample. This approach
is particularly popular for the
quantitation of endogenous proteins
such as biomarkers, for which
well‑characterized immunological
reagents are widely available.
One example is an LC–MS–MS
method for parathyroid hormone
(PTH) in human serum (9). A
sample of 1 mL was treated by IC
with polystyrene beads coated
with murine anti‑PTH antibodies
and the trapped analyte digested
with trypsin. The IC treatment
allowed the quantitation of PTH
down to 40 pg/mL (4 pM) in serum,
which shows the enormous clean‑
up potential of this approach. A
completely 15N‑labelled form of
PTH was added to the sample as
40 Advances in Biopharmaceutical Analysis – October 2015
van de Merbel
2.94
1000
800
600
400
200
Inte
nsi
ty, cp
sIn
ten
sity
, cp
s
0
1000(b)
(a)
800
600
400
200
0
0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0
0.5 1.0 1.5 2.0
Time (min)
Time (min)
2.5 3.0 3.5 4.0 4.5 5.0
1.04
1.04
2.96
3.20
3.43 4.054.12
1.441.52 1.62 2.50 2.64
3.173.40
3.49 3.69
4.01
4.11
Figure 2: LC–MS–MS (m/z 616.1 to m/z 1030.7) chromatogram of the signature peptide of recombinant human alpha‑glucosidase in human plasma; (a) blank and (b) 0.5 µg/mL, pretreated with digestion only.
100 2.58e3
6.63
6.51
6.65
6.78
6.94
7.007.13
7.237.49
7.55
7.737.91
6.71
6.79 6.92
7.097.507.55
7.63
7.817.90
6.60 6.80 7.00 7.20 7.40 7.60 7.80
[rhTRAIL]=10 ng/mL [rhTRAIL]=10 ng/mL
SCX
0
%
100 2.02e3
6.60 6.80 7.00 7.20 7.40 7.60 7.80
Time (min)Time (min)
IMAC
0
%
Figure 3: LC–MS–MS (m/z 729.0 to m/z 942.4) chromatograms of the signature peptide of 10 ng/mL rhTRAIL in human serum and the corresponding blanks, pretreated with SCX or IMAC before digestion. Adapted and reproduced with permission from Bioanalysis 7(6), D.Wilffert, R. Bischoff, and N.C. van de Merbel, Antibody-free workflows for protein quantification by LC-MS/MS, 763–779 (2015) © Future Science Ltd.
ES683325_LCESUPP1015_040.pgs 10.01.2015 20:52 ADV blackyellowmagentacyan
an internal standard at the very
beginning of the sample handling
procedure. In general, it is desirable
that a stable‑isotope‑labelled or
other closely related form of the
protein analyte be included in the
method as an internal standard,
to correct for the variability of the
extraction procedure. This is also
one of the drawbacks of extracting a
biological sample before digestion,
because such a protein‑based
internal standard can usually only
be obtained by biotechnological
means, which may be difficult, if not
impossible (4).
The disadvantages associated with
the use of immunological reagents
— such as their potentially limited
availability, varying quality, and
the interference of matrix proteins
with the extraction efficiency —
have prompted researchers to
investigate alternative so‑called
antibody‑free extraction approaches
(5). An interesting technique
is immobilized‑metal affinity
chromatography (IMAC), which is
based on the interaction of metal
ions, such as Ni2+, with amino acids
that feature strong electron donor
groups, such as histidine. Proteins
with such amino acids on their
surface will be selectively captured
by IMAC resins. As an example,
the biopharmaceutical recombinant
human tumour necrosis factor‑related
apoptosis‑inducing ligand (rhTRAIL)
has been quantified in human and
mouse serum down to 20 ng/mL
(340 pM) by removing 95% of matrix
proteins, while recovering >70% of
the analyte with IMAC (8).
Another technique is solid‑phase
extraction (SPE) with ion‑exchange
materials, which separates proteins
based on their isoelectric point (pI).
Proteins with a relatively high pI
bear a net positive charge and can
be trapped on a cation‑exchange
resin at neutral or slightly alkaline
pH, at which many endogenous
proteins with a lower pI will be
negatively charged and thus not be
captured. The extraction of rhTRAIL
with strong‑cation exchange (SCX)
SPE was found to have a similar
clean‑up potential to IMAC, with
an analyte recovery of 70% and a
protein removal efficiency of 99%.
As an illustration, Figure 3 shows
chromatograms obtained for 10 ng/
mL (170 pM) of rhTRAIL in human
serum, which was extracted by
SCX or IMAC, followed by trypsin
digestion and LC–MS–MS analysis of
the signature peptide.
Peptide ExtractionRemoval of interfering matrix
components is also possible
after digestion, that is, at the
peptide level. This approach
has some distinct advantages.
From a practical point of view the
optimization of an SPE procedure
is more straightforward because of
the wide availability of a range of
materials that are commonly used
for small‑molecule extractions and
41www.chromatographyonline.com
van de Merbel
100
%
0
100
%
0
2.32
1.51
3.90
4.275.07 5.25
5.715.84
4.59
2.843.52
4.06
4.204.55
5.03
5.30
5.47
6.19
5.88
(a)
(b)
Time (min)
Time (min)
Figure 4: LC–MS–MS (m/z 752.0 to m/z 773.3) chromatograms of the signature peptide of 10 ng/mL of a nanobody in human plasma (a) without or (b) with solid‑phase extraction of the plasma digest. Analyte peak at 4.6 min. Adapted and reproduced with permission from Bioanalysis 7(1), K.J. Bronsema, R. Bischoff, M.P. Bouche, K. Mortier, and N.C. van de Merbel, High-sensitivity quantitation of a Nanobody® in plasma by single-cartridge multidimensional SPE and ultra-performance LC-MS/MS, 53–64 (2015) © Future Science Ltd.
100
0
12.50 13.00
12.70
(a)
Time (min) Time (min)
(b)
%
100
0
13.00 14.00 15.00
15.4414.72
13.52
12.76
%
Figure 5: LC–MS–MS (m/z 456.6 to m/z 852.5) chromatograms of the signature peptide (a) of 0.2 ng/mL of rhTRAIL spiked to dog saliva, plus corresponding blank, or (b) of endogenous human TRAIL in unspiked human saliva. Samples pretreated with IMAC before and SCX after digestion.
ES683322_LCESUPP1015_041.pgs 10.01.2015 20:53 ADV blackyellowmagentacyan
because of the more predictable
extraction behaviour of smaller
peptides compared to that of intact
proteins.
The accuracy and precision of
extractions may be influenced
by protein‑protein interactions in
samples (such as binding of a
biopharmaceutical to its target
or to anti‑drug antibodies), or the
occurrence of aggregates. If a
sample is first subjected to digestion,
these interactions will no longer
influence the extraction because all
proteins will have been cleaved to
peptides that are much less likely
to bind to one another with a high
affinity.
No less importantly, peptide
extraction does not need a
protein‑based internal standard; it
can instead perform very well when
using a stable‑isotope labelled form
of the signature peptide (4,6), which
is considerably less expensive and
easier to obtain. It may, however,
be difficult to achieve sufficient
selectivity because the peptides
in a plasma digest are much more
similar to each other than the plasma
proteins were before digestion.
Again, the highest selectivity and
sensitivity is achieved by applying
immunocapture, which in this case
uses immobilized antibodies raised
against the signature peptide. This
approach is most widespread in the
field of biomarker analysis, where
the number of analytes is relatively
limited and assays are relevant to
many research groups around the
globe. Large clean‑up efficiencies
can be achieved in this way, as
was reported for the endogenous
proteins α1‑antichymotrypsin
(1453‑fold enrichment relative to
matrix proteins) and TNF‑α (573‑fold
enrichment) (10).
IC at the peptide level is less
popular in biopharmaceutical
analysis, probably because
of the general drawbacks of
antibody‑based reagents with regard
to availability and batch‑to‑batch
reproducibility. A more generic
approach for peptide extraction
from a digest is to use conventional
ion‑exchange SPE, but this needs
to be carefully optimized to obtain
sufficient selectivity. A digest of a
protein‑rich biological sample (such
as plasma) contains a multitude of
peptides, which all have carboxylic
and amine groups, and the signature
peptide can only be separated
from the excess of endogenous
background peptides if its pI value
is sufficiently different. Typically, the
pH and ionic strength of the loading,
washing, and elution steps need to
be carefully optimized for a selective
extraction.
A biopharmaceutical nanobody
was quantified down to 10 ng/
mL (360 pM) in rabbit and human
plasma by trypsin digestion followed
by SPE on a weak‑anion exchange
phase (11). The signature peptide
contained three carboxylic acid
groups and was strongly retained
by the positively charged SPE
phase at pH 5; many endogenous
peptides with less negative charges
were not trapped during sample
loading or were removed from the
SPE material by a washing step
with 300 mM sodium chloride. The
mixed‑mode SPE phase, which also
contained reversed‑phase groups,
was subsequently neutralized at a
high pH and the (relatively polar)
signature peptide was eluted,
while some less polar endogenous
peptides remained bound by
reversed‑phase interactions. In this
way, two dimensions of selectivity
(ion exchange and reversed phase)
were used to isolate the signature
peptide from the plasma digest.
Figure 4 illustrates that many
interfering peaks were removed
from the chromatogram with this
approach and that selectivity was
clearly improved. Of course, cation‑
exchange SPE can be applied in
the same way in case the signature
peptide has multiple positive
charges, and even reversed‑phase
SPE might be an option if the
signature peptide is particularly
hydrophobic.
Combined Protein and Peptide ExtractionAs illustrated above, generic protein
or peptide extractions typically result
in LLOQs in the low ng/mL (mid to
high pM) range, while IC extraction at
42 Advances in Biopharmaceutical Analysis – October 2015
van de Merbel
400
350
300
250
200
150
100
50
QLI
DIV
DQ
LK In
ten
sity
(a) (b)
04.5 5.0
Time (min) Time (min)
5.5
400
350
300
250
200
150
100
50
04.5 5.0 5.5
Figure 6: LC–MS–MS chromatograms of two mass transitions (m/z 592.8 to m/z 943.5 in red and m/z 592.8 to m/z 830.5 in blue) of the signature peptide of IL‑21 in human serum; (a) blank and (b) 0.78 pg/mL. Samples pretreated with immunocapture both before and after digestion. Adapted and reproduced with permission from Analytical Chemistry 85, J. Palandra, A. Finelli, M. Zhu, et al., Highly Specific and Sensitive Measurements of Human and Monkey Interleukin 21 Using Sequential Protein and Tryptic Peptide Immunoaffinity LC–MS/MS, 5522–5529 (2013) © American Chemical Society.
ES683324_LCESUPP1015_042.pgs 10.01.2015 20:53 ADV blackyellowmagentacyan
Advances in Biopharmaceutical Analysis – OCTOBER 2015 43
ADVERTISEMENT FEATURE
Gel permeation chromatography (GPC), also known as
size-exclusion chromatography (SEC), provides an easy
and effective way to measure the molar mass distribution
and the amount of free, unbound polysaccharide in iron
polysaccharide complexes.
Iron is an essential nutrient in the human body. In case of iron deficiency, complexes of a polysaccharide and iron are applied as drugs to enhance low iron levels. Suitable characterization of these complexes and their formulations are mandatory for regulatory reasons, quality control, and research. In the present investigation, iron polysaccharide complexes from different sources were analyzed on a GPC/SEC system with simultaneous ultraviolet/refractive index (UV/RI) detection.
Experimental Conditions:
GPC/SEC was performed using a PSS BioSECcurity SEC systemColumns: PSS SUPREMA, 5 µm, 30 Å + 2 ×1000 Å (8 × 300 mm, each) PSS SUPREMA precolumnEluent: 0.1 n NaNO3, in 0.01 m phosphate buffer at pH = 7Temperature: AmbientDetection: UV @ 254 nm, refractive index (RI)Calibration: PSS Pullulan ReadyCal standards Concentration: 2 g/L for dry material, approx. 50 g/L for formulationsInjection volume: 25 µLSoftware: PSS WinGPC UniChrom 8.2
Results and Discussion:
Figure 1 shows the overlay of the UV-chromatograms of the four different samples A, B, C, and D, while the inset of the figure shows the overlay of the simultaneously measured RI-traces for two of the samples (A and B), which show nearly identical UV-traces.
An advantage of this application is that the iron polysaccharide complex is selectively detected by the UV-detector operated at 254 nm (20–26 mL). All complexes reveal well shaped nearly Gaussian peak shapes, indicating that the PSS SUPREMA column combination is ideal for this molar mass separation range. By applying a calibration curve, established using PSS pullulan standards, the relative molar mass distributions as well as the molar mass averages and the polydispersities are derived.
While UV-detection is sufficient to differentiate between three of the four samples, samples A and B render identical elution profiles. However, when comparing the RI-traces of both samples, it becomes clear that sample A contains a significantly higher amount of the unbound polysaccharide.
We can therefore conclude that GPC/SEC with UV- and RI-detection does not only allow the molar mass distribution of iron polysaccharide complexes to be determined, but also provides information on the amount of free, unbound polysaccharide ensuring a more comprehensive characterization of the samples.
Investigation of Iron Polysaccharide Complexes by GPC/SEC Using RI- and UV-DetectionPSS Polymer Standards Service GmbH
PSS Polymer Standards Service GmbHIn der Dalheimer Wiese 5, D-55120 Mainz, Germany
Tel: +49 6131 962390 fax: +49 6131 9623911
E-mail: [email protected]
Website: www.pss-polymer.com
Figure 1: Comparison of the UV-traces of four different iron dextran samples used to determine the molar mass distribution of the iron complexes. While the UV-signals for samples A and B are nearly identical, the inset displaying the RI-traces shows that these samples differ in the amount of unbound polysaccharide.
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the protein or peptide level enables
quantitation down to mid pg/mL (low
to mid pM) concentrations. If more
sensitivity is required, one option is
to combine protein and peptide
extractions. Excellent selectivity
and sensitivity can be reached even
without antibody‑based extraction
materials, as was shown for rhTRAIL
in saliva (12). After IMAC extraction
of the protein analyte and trypsin
digestion, the digest was further
purified using SPE on a SCX
cartridge. Because of the presence
of four basic amino acids in the
signature peptide, the digest was
acidified before loading onto the SPE
phase. The peptide was then trapped
and endogenous peptides were
removed by washing with 200 mM
sodium chloride. After elution at
alkaline pH, the signature peptide
was quantified using LC–MS–MS.
As shown in Figure 5, a TRAIL
concentration as low as 0.2 ng/mL
(3.4 pM) could be quantified in both
dog and human saliva. In principle,
protein or peptide extractions can
be combined in many ways and as
long as the separation mechanisms
are orthogonal, improved selectivity
and sensitivity can be expected
compared to a single‑extraction
approach.
The ultimate combination of
protein and peptide extraction is IC
of the protein analyte followed by
digestion and IC of the signature
peptide. Although this requires two
specifically raised antibodies and is
by no means a generic approach, it
can result in impressive sensitivities.
The biomarker interleukin‑21 (IL‑21)
was quantified in human serum
and monkey tissues with an LLOQ
of 0.78 pg/mL (0.05 pM). This was
achieved by combining off‑line
magnetic bead‑based protein
extraction using an anti‑IL‑21
antibody with on‑line enrichment
of the signature peptides using
immobilized anti‑peptide antibodies
(13). Figure 6 shows representative
chromatograms. It is important
to realize that the obtained LLOQ
corresponds to a molar concentration
of the protein, which is five orders
of magnitude lower than that shown
in Figure 2 (digestion only). This
convincingly demonstrates the
enormous clean‑up capability of this
combination of techniques.
LC–MS–MS versus ELISACompared to ligand‑binding
assays, LC–MS–MS has a number
of analytical advantages such as
a larger linear dynamic range;
(usually) higher accuracy and
precision because of the possibility
to apply internal standards (4); the
ability to quantify multiple analytes
simultaneously; and the fact that
it does not necessarily require
immunological reagents (5). The
last point can be especially critical,
because such reagents may be
problematic to obtain or show a
large batch‑to‑batch variability,
which makes comparison of results
between laboratories or over a
longer period of time difficult, if not
impossible. The disadvantages of
LC–MS–MS include its generally
higher operational cost; more
limited sample throughput; and less
favourable concentration sensitivity.
In addition, with the digestion step
that is generally needed for LC–MS–
MS, the three‑dimensional structure
of a protein analyte is lost and the
analytical principle is therefore not
related to the complex molecular
structure of a protein, which
determines its pharmacological
activity.
Now that more and more reports
are appearing that compare newly
developed LC–MS–MS methods
with existing ELISAs for the same
protein analyte, it is becoming
increasingly clear that both
techniques do not always give
superimposable concentration
results (14,15). Although in the world
of small‑molecule quantitation,
two different results for the same
sample would be seen as proof that
at least one of them is incorrect,
this is not necessarily true for
biopharmaceuticals. It should be
realized that, in contrast to small
molecules, LC–MS–MS as well
as ELISA only use a small part
of the protein molecule for the
actual quantitation, the signature
peptide and the binding epitope,
respectively, and this may represent
as little as a few percent of the
entire molecule. Furthermore,
both techniques are based on
quite different (bio)chemical
principles, to which the structurally
complex and often heterogeneous
biopharmaceuticals may respond
in different ways. Thus, neither
LC–MS–MS nor ELISA should be
regarded as the ultimate quantitation
technique for biopharmaceuticals,
but rather as complementary tools
for obtaining quantitative information
about this complicated but very
interesting class of compounds.
AcknowledgementStichting Technische Wetenschappen
(STW) and Samenwerkingsverband
Noord‑Nederland (SNN) are
gratefully acknowledged for
providing financial support for part of
the work described in this paper.
References(1) R. Bischoff, K.J. Bronsema, and N.C.
van de Merbel, Trends Anal. Chem. 48,
41–51 (2013).
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(2013).
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894, 1–14 (2012).
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(11) K.J. Bronsema, R. Bischoff, M.P.
Bouche, K. Mortier, and N.C. van de
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(12) D. Wilffert, unpublished results
(13) J. Palandra, A. Finelli, M. Zhu, J.
Masferrer, and H. Neubert, Anal. Chem.
85, 5522–5529 (2013).
(14) N.C. van de Merbel, K.J. Bronsema,
and M. Nemansky, Bioanalysis 4, 2113–
2116 (2012).
(15) P. Bults, N.C. van de Merbel, and R.
Bischoff, Expert Rev. Proteomics 12,
355–374 (2015).
Nico van de Merbel is scientific
director at the bioanalytical
laboratories of PRA Health Sciences
(Assen, The Netherlands and
Lenexa, KS, USA) and honorary
professor at the University of the
Groningen, The Netherlands.
44 Advances in Biopharmaceutical Analysis – October 2015
van de Merbel
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45www.chromatographyonline.com
Ph
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In contrast to small molecule drugs
that are commonly synthesized
by chemical means, protein
biopharmaceuticals result from
recombinant expression in
non‑human host cells. As a result,
the biotherapeutic is co‑expressed
with hundreds of host cell proteins
with different physicochemical
properties present in a wide dynamic
concentration range. During
downstream processing, the levels
of HCPs are substantially reduced
to a point considered acceptable
to regulatory authorities (typically
< 100 ppm–ng HCP/mg product).
These process‑related impurities
are considered as critical quality
attributes because they might induce
an immune response, cause adjuvant
activity, exert a direct biological
activity (such as cytokines), or act on
the therapeutic itself (for example,
proteases) (1,2). To mention some
specific examples, during the clinical
development phase of Omnitrope,
Sandoz’s human growth hormone
biosimilar expressed in E. coli,
adverse events associated with
residual HCPs were encountered.
The European Medicines Agency
(EMA) only granted approval after
additional purification steps for
HCP clearance were incorporated
(3–5). Scientists at Biogen Idec
demonstrated fragmentation of a
highly purified monoclonal antibody
as a result of residual Chinese
Hamster Ovarian (CHO) cell protease
activity in the drug substance,
despite an enormous purification
effort undertaken (Protein A affinity
chromatography with subsequent
orthogonal purification steps
by cation‑ and anion‑exchange
chromatography) (6). The authors
of the study state that it is of utmost
importance to identify residual
protease activity early in process
development to allow a revision of
the purification scheme or ultimately
to knockdown the specific protease
gene.
Multicomponent enzyme‑linked
immunosorbent assay (ELISA) is
presently the workhorse method
for HCP testing because of its high
throughput, sensitivity, and selectivity
(1,2). Polyclonal antibodies used in
the test are typically generated by
the immunization of animals with
an appropriate preparation derived
from the production cell, minus the
product‑coding gene. However,
ELISA does not comprehensively
recognize all HCP species, that
is, it cannot detect HCPs to which
no antibody was raised, it only
provides information on the total
amount of HCPs without providing
insight in individual HCPs, and, in
a multicomponent set‑up, it has
a poor quantitation power. In that
respect, MS nicely complements
ELISA because it can provide
both qualitative and quantitative
information on individual HCPs. In
recent years, various papers have
appeared dealing with the mass
spectrometric analysis of HCPs
(2,3,7–12). These studies typically
rely on bottom‑up proteomics
approaches in which peptides
derived from the protein following
proteolytic digestion are handled.
A clear trend is observed towards
the use of upfront multidimensional
chromatography to tackle the
enormous complexity and wide
dynamic range (2,3,7,8,10).
Compared to one‑dimensional
LC (1D‑LC), two‑dimensional LC
(2D‑LC) drastically increases
peak capacity as long as the two
dimensions are orthogonal (13). In a
one‑dimensional chromatographic
set‑up the separation space is
dominated by peptides derived from
the therapeutic protein, in 2D‑LC
the increased peak capacity allows
one to look substantially beyond
the therapeutic peptides and detect
Analyzing Host Cell Proteins Using Off‑Line Two-Dimensional Liquid Chromatography–Mass SpectrometryKoen Sandra, Alexia Ortiz, and Pat Sandra, Research Institute for Chromatography (RIC) and Metablys, Kortrijk, Belgium.
Protein biopharmaceuticals are commonly produced recombinantly in mammalian, yeast, or bacterial expression systems. In addition to the therapeutic protein, these cells also produce endogenous host cell proteins (HCPs) that can contaminate the biopharmaceutical product, despite major purifcation efforts. Since HCPs can affect product safety and efficacy, they need to be closely monitored. Enzyme-linked immunosorbent assays (ELISA) are recognized as the gold standard for measuring HCPs because of their high sensitivity and high throughput, but mass spectrometry (MS) is gaining acceptance as an alternative and complementary technology for HCP characterization. This article reports on the use of off-line two-dimensional liquid chromatography–mass spectrometry (2D-LC–MS) for the characterization of HCPs and their monitoring during downstream processing.
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46 Advances in Biopharmaceutical Analysis – October 2015
Sandra et al.
HCPs at low levels. Three recent
papers using 2D‑LC–MS–MS
demonstrate that HCPs can be
revealed at levels as low as 10 ppm
(2,3,7). In these cases, label‑free
quantification was based on the three
most intense tryptic peptides making
use of single‑point calibration against
spiked exogenous proteins.
An off‑line 2D‑LC–MS–MS setup
was used in our laboratory for the
characterization of HCPs throughout
the downstream manufacturing of a
therapeutic enzyme recombinantly
expressed in yeast. The workflow is
schematically presented in Figure 1.
Supernatant was collected at different
purification steps. Following desalting
of the supernatant, the proteins were
reduced using dithiothreitol (DTT)
and alkylated using iodoacetamide
(IAM) prior to overnight trypsin
digestion. The peptide mixture was
subsequently subjected to 2D‑LC–
MS–MS.
In successfully applying 2D‑LC,
the selectivity of the two separation
mechanisms towards the peptides
must differ substantially in order to
maximize orthogonality and, hence,
resolution. Various orthogonal
combinations targeting different
physicochemical properties of the
peptides have been described.
Bottom‑up proteomics set‑ups
initially relied on the combination of
strong‑cation exchange (SCX) and
reversed‑phase LC to separate by
charge in the first dimension and
by hydrophobicity in the second
dimension (13–15). In recent years,
various researchers have shifted
their efforts to the combination
of reversed‑phase LC and
reversed‑phase LC (13,16–19).
The orthogonality in this non‑
obvious combination is mainly
directed by the mobile phase pH,
in this instance, high pH in the
first dimension and low pH in the
second dimension, and by the
zwitterionic nature of the peptides.
In contrast to the combination
of SCX and reversed‑phase LC,
where the first dimension has an
intrinsic low peak capacity, the
combination of reversed‑phase LC
in both dimensions benefits from
the high peak capacities of the two
independent dimensions, which
results in an overall high peak
capacity of the 2D set‑up.
Supernatants
Buffer exchange/desalting
Reduction/alkylation/digestion
Database search
Protein ID/Quant
High pH reversed-phase LC x low pH reversed-phase LC
QTOF MS–MS
Figure 1: Workflow for the characterization of HCPs using off‑line 2D‑LC–MS–MS.
Time (min)
0 5 10 15 20 25 30 35
mA
U
0
200
400
600
800
1000
1200
1400
Fraction 11-19
Figure 2: First‑dimension reversed‑phase LC–UV 214 nm chromatogram of a selected downstream manufacturing sample. HPLC system: Agilent Technologies 1200; Column: 2.1 mm × 150 mm, 3.5‑µm Waters XBridge BEH C18; Mobile phase A: 10 mM NH4HCO3 pH 10; Mobile phase B: acetonitrile; Flow rate: 200 µL/min; Gradient: 5–50% B in 30 min; Column temperature: 25 °C; Injection volume: 50 µL; Fraction interval: 1.5 min (300 µL fractions).
ES683398_LCESUPP1015_046.pgs 10.01.2015 21:03 ADV blackyellowmagentacyan
47www.chromatographyonline.com
Sandra et al.
the final stages of purification. Of
particular interest, during downstream
manufacturing, a non‑yeast derived
glycosidase was added to shape the
glycosylation profile of the therapeutic
enzyme (in between stage 1 and 2).
This glycosidase temporarily reduced
the purity of the therapeutic enzyme
but was rapidly cleared. The HCPs
detected were mainly proteases,
which influenced stability of the
therapeutic enzyme. While some
were clearly reduced throughout the
process (serine carboxypeptidase 1
and aspartyl peptidase), others were
enriched (serine carboxypeptidase 2
and metallopeptidase). While these
proteases were present at low levels
(<0.1%), stability studies have shown
that they act on the protein. With the
identity of these proteases revealed,
they could be the subject of a gene
knockout to increase product stability.
It is important to note that none of
the HCPs reported could be identified
using 1D‑LC–MS–MS operated
under exactly the same conditions
as reported in the legend of Figure
In the characterization of
yeast HCPs, we opted to use
reversed‑phase LC in both
dimensions with the first dimension
operated at pH 10 and the second
dimension at pH 2.6. An acidic
pH is preferred in the second
dimension since it maximizes MS
sensitivity for peptides. Figure
2 shows the first dimension UV
214‑nm chromatogram of a selected
downstream manufacturing sample.
A reversed‑phase LC column with an
internal diameter of 2.1 mm was used,
which allowed substantial amounts of
sample to be loaded, in this particular
case the amount corresponding
to 115 µg of protein. The peptides
were nicely spread throughout the
acetonitrile gradient and 22 fractions
were collected and further processed
after drying and reconstitution in
50 µL low pH mobile phase A (2%
acetonitrile and 0.1% formic acid).
The second dimension consisted
of a reversed‑phase LC capillary
column with an internal diameter of
75 µm, which was directly coupled
through a nanospray interface to high
resolution quadrupole time‑of‑flight
(QTOF‑) MS operated in the data‑
dependent acquisition (DDA) mode.
The LC–MS–MS traces of some
selected fractions are shown in Figure
3 illustrating good orthogonality
between first and second dimension
separations.
The MS system was programmed
so that an MS survey measurement
preceded three dependent MS–
MS acquisitions. Precursors
selected twice for collision‑induced
dissociation (CID) were placed
in an exclusion list. Generated
MS–MS spectra were subjected to
database searching (yeast proteins
and therapeutic enzyme sequence)
and relative protein quantification
was performed from total protein
intensities computed by the Spectrum
Mill search engine. Total intensity is
the sum of intensities for all spectra
of peptides belonging to a given
protein. Figure 4 shows the evolution
of the therapeutic enzyme and
some selected HCPs throughout
7x10
0
7x10
0
7x10
0
7x10
0
7x10
0
7x10
0
7x10
0
7x10
0
7x10
0
MS counts vs. Acquisition Time (min)
4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 34 36 38 40
Fraction 12
Fraction 13
Fraction 14
Fraction 15
Fraction 16
Fraction 17
Fraction 18
Fraction 19
Fraction 11
Figure 3: Second dimension LC–MS–MS chromatograms of selected fractions (Figure 2). HPLC system: ThermoScientific Ultimate3000 RSLC nano; MS system: Agilent Technologies 6530 Q‑TOF, Column: 75 µm × 150 mm, 3‑µm ThermoScientific Acclaim PepMap100 C18, Pre‑column: 75 µm × 20 mm, 3‑µm Acclaim PepMap100 C18 (Thermo Scientific), Mobile phase A: 2% acetonitrile, 0.1% formic acid, Mobile phase B: 80% acetonitrile, 0.1% formic acid; Loading solvent: 2% acetonitrile, 0.1% formic acid; Flow rate: 300 nL/min (nano pump), 5 µL/min (loading pump); Gradient: 0–60% B in 60 min; Column temperature: 35°C; Injection volume: 20 µL.
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48 Advances in Biopharmaceutical Analysis – October 2015
Sandra et al.
3. Column load was evidently much
lower compared to the 2D‑LC–MS–
MS analysis (4 µg vs. 115 µg).
In conclusion, off‑line 2D‑LC–
MS–MS represents a valuable new
tool for the characterization of HCPs
and their monitoring throughout
downstream processing. The use of
multidimensional chromatography
substantially increases peak capacity
and improves the dynamic range
providing access to otherwise
unmined HCPs. Based on the output
of the 2D‑LC–MS–MS experiment,
processes can be adjusted and
identified HCPs can be incorporated
in single product ELISAs or in
targeted multiple reaction monitoring
(MRM) MS assays for routine
monitoring.
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Koen Sandra is Director
at the Research Institute for
Chromatography (RIC, Kortrijk,
Belgium).
Alexia Ortiz is a Proteomics
Researcher at the Research Institute
for Chromatography (RIC, Kortrijk,
Belgium).
Pat Sandra is Chairman at
the Research Institute for
Chromatography (RIC, Kortrijk,
Belgium) and Emeritus Professor at
Ghent University (Ghent, Belgium).
Recombinant therapeutic enzyme
99.53
45
99.47
46
0.44
0.008
0.0130.43
0.015
7 1
211
20.000.07
5
0.054
0.023
99.77
0.0346
3
0.06
3
0.09
9
Exogenous glycosidase Metallopeptidase (HCP)
Serine carboxypeptidase 1 (HCP) Aspartyl peptidase (HCP)
Pe
rce
nta
ge
Pe
rce
nta
ge
Pe
rce
nta
ge
Pe
rce
nta
ge
Pe
rce
nta
ge
Pe
rce
nta
ge
Purifcation stage
1 2 3 1 2 3
Purifcation stage
Purifcation stage
1 2 3 1 2 3
Purifcation stage
Purifcation stage
1 2 3 1 2 3
Purifcation stage
Serine carboxy peptidase 2 (HCP)
Figure 4: Evolution of the therapeutic enzyme, the exogenous glycosidase, and some selected HCPs throughout the final stages of downstream manufacturing. The numbers on the bars represent the relative abundances and the number of unique peptides identified and quantified. Relative abundances were calculated based on the MS signal of identified peptides. Note: the therapeutic enzyme contains various fully occupied glycosylation sites. These glycopeptides are not identified by the MS–MS search engine and therefore not taken into account in the calculation of relative abundances.
ES683394_LCESUPP1015_048.pgs 10.01.2015 21:03 ADV blackyellowmagentacyan
Advances in Biopharmaceutical Analysis – OCTOBER 2015 49
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limits. To simplify this process, Shimadzu has devised a novel
technique — nSMOL (nano-surface and molecular-orientation
limited proteolysis) (Figure 1) — that can be applied to all mAbs.
Experimental Workflow
nSMOL works on selective proteolysis of Fab by making use
of the difference in size of the protease nanoparticle diameter
(200 nm) and the antibody resin pore size (100 nm) (Figure 2).
In a first step, plasma applied on the resin allows the antibody
Fc to bind to the protein A/G inside the pore of the resin. After
washing steps, the protease on the nanoparticle surface is
applied to the resin to proteolyze only the Fab region, which
leads to specific CDR peptide collection after filtration. CDR
peptides can then be quantified on the LC–MS–MS system.
By using nSMOL, one can maintain the specificity of the
antibody sequences while minimizing sample complexity as
well as eliminating extra protease. This approach can lead to
a shortened analytical time, increased LC–MS–MS robustness,
a wide dynamic range, and considerable improvement in
sensitivity.
This technique can already boast of being ready to use
with completely validated methodology for the bioanalysis of
Trastuzumab and other antibody drugs in human plasma in
accordance with the Japan Guideline on Bioanalytical Method
Validation in Pharmaceutical Development from Notification 0711-
1(2013) of the Evaluation and Licensing Division, Pharmaceutical
and Food Safety Bureau, the Ministry of Health, Labour and
Welfare, dated 11 July 2013.
Changes Everything — the New LCMS
To address the second challenge, Shimadzu has introduced the
new LCMS-8060 triple quadrupole mass spectrometer (Figure
3), which is part of the Ultra-fast Mass Spectrometry (UFMS)
platform of MS–MS systems. With a new UF Qarray ion guide
technology increasing ion production and signal intensity while
maintaining very low background noise, the LCMS-8060 brings
a new distinct vision of sensitivity that makes a real difference
in working better and faster. In short, LCMS-8060 is designed
nSMOL: Limited Proteolysis on the Fab and Accelerating mAb Bioanalysis Using LC–MS–MSDr Takashi Shimada1 and Stéphane Moreau2, 1Shimadzu Corporation, 2Shimadzu Europa GmbH
Figure 1: nSMOL technology.
Figure 2: Resin with protein A/G inside the pore and nanoparticles.
ES683353_LCESUPP1015_049.pgs 10.01.2015 20:55 ADV blackyellowmagentacyan
50 Advances in Biopharmaceutical Analysis – OCTOBER 2015
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to push the limits of quantitation for complex bioanalysis
experiments by providing the highest sensitivity, durability, and
a short analysis time.
Conclusion
Only when powerful techniques complement each other can
scientists find a whole solution to their challenges. nSMOL
technology with the LCMS-8060 combines perfectly to
solve biosimilar and mAb quantitation puzzles with the right
perspective during both pre-clinical and clinical phases of
development.
Reference
(1) Noriko Iwamoto, Takashi Shimada, Yukari Umino, et al., Analyst 139,
576–580 (2014).
(2) Noriko Iwamoto, Yukari Umino, Takashi Shimada, et al., Anal. Methods
DOI:10.1039/C5AY01588J (2015).
Shimadzu Europa GmbHAlbert-Hahn-Str. 6–10, D-47269 Duisburg, Germany
Tel: +49 203 76 87 0 fax: +49 203 76 66 25
E-mail: [email protected]
Website: www.shimadzu.eu
Figure 3: LCMS-8060 Triple Quadrupole Mass Spectrometer.
ES683354_LCESUPP1015_050.pgs 10.01.2015 20:55 ADV blackyellowmagentacyan
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