Advances in Biopharmaceutical Analysisfiles.alfresco.mjh.group/alfresco_images/pharma/... · Chief Financial Officer Robert Gray Chairman Dame Helen Alexander CORPORATE OFFICE 641

Supplement tooctober 2015

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Biopharmaceutical Analysis

Advances in

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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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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.


Pat Sandra Koen Sandra


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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.

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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


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.


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


AdvanceBio

RP‑mAbC8, C4, diphenyl

3.5 (0.25‑µm


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


Flare Widepore (Diamond Analytics) C183.6 (0.1‑µm


ES683260_LCESUPP1015_010.pgs 10.01.2015 20:46 ADV blackmagentacyan


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


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.


(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


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.


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


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.


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).

References(1) C.J.V. Oss, R.J. Good, and M.K. Chaudhury,

J. Chromatogr. 376, 111–119 (1986).(2) J.A. Querioz, C.T. Tomaz, and J.M.S.

Cabral, J. Biotech. 87, 143–159 (2001).(3) L.N. Lee, J.M.R. Moore, J. Ouyang, X.

Chen, M.D.H. Nguyen, and W.J. Galush, Anal. Chem. 84, 7479–7486 (2013).

(4) Lan N. Le, Jamie M.R. Moore, Jun Ouyang, et al., Analytical Chemistry 84, 7479–7486 (2012).

(5) F. Debaene, A. Boeuf, E. Wagner‑Rousset, O. Colas, D. Ayoub, N. Corvaïa, A. Van Dorsselaer, A. Beck, and S. Cianférani, Anal. Chem. 86, 10674–10683 (2001).

(6) M. Haverick, S. Mengisen, M. Shameem, and A. Ambrogelly, mAbs 6, 852–858 (2001).

(7) S. Fekete, A. Beck, and D. Guillarme, Am.

Pharm. Rev. 18, 59–63 (2015).(8) S. Fekete, A. Beck, and D. Guillarme,

J. Pharm. Biomed. Anal. 111, 169–176 (2015).

(9) Szabolcs Fekete, Alain Beck, and Davy Guillarme, Journal of Pharmaceutical and

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,

Anal. Chem. 39, 1422–1428 (1967).(19) C. Horvath and S.R. Lipsky, J.

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,

and J.J. Kirkland, J. Chromatogr. A 1315, 118–126 (2013).

(22) S. Fekete, K. Ganzler, and J. Fekete, J. Pharm. Biomed. Anal. 54, 482–490 (2011).

(23) F. Gritti and G. Guiochon, J. Chromatogr.

A 1218, 907–921 (2011).(24) S.A. Schuster, B.M. Wagner, B.E. Boyes,

and J.J. Kirkland, J. Chromatogr. Sci. 48, 566–571 (2010).

(25) S. Fekete and D. Guillarme, J.

Chromatogr. A 1320, 86–95 (2013). (26) B. Bobály, D. Guillarme, and S. Fekete, J.

Sep. Sci. 37, 189–197 (2014). (27) B. Bobály, D. Guillarme, and S. Fekete,


(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

1270, 194–203 (2012).(30) L.E. Blue and J.W. Jorgenson, J.

Chromatogr. A 1218, 7989–7995 (2011).(31) C. Viklund, F. Svec, J.M.J. Fréchet, and K.

Irgum, Chem. Mater. 8, 744–750 (1996).(32) M. Petro, F. Svec, I. Gitsov, and J.M.J.

Fréchet, Anal. Chem. 68, 315–321 (1996). (33) S. Fekete, J.‑L. Veuthey, S. Eeltink, and

D. Guillarme, Anal. Bioanal. Chem. 405, 3137–3151 (2013).

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.


Guillarme et al.


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.



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


Sandra et al.

0,8

0,7

0,6

0,5

0,4

0,3

0,2

0,1

0,03

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600

500

400

300

200

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25 26 27 28 32

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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.


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


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.


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


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


22800 23000 23200 23400 23600 23800 24000 24200 24400


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.


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


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.


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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


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.


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).

References

(1) K. Strebhardt and A. Ullrich, Nat. Rev.

Cancer 8, 473–480 (2008).

(2) L.M. Weiner, R. Surana, and S. Wang,

Nat. Rev. Immunol. 10, 317–327 (2010).

(3) G. Köhler and C. Milstein, Nature 256,

495–497 (1975).

(4) N.A.P.S. Buss, S.J. Henderson, M.

McFarlane, J.M. Shenton, and L. de

Haan, Curr. Opin. Pharmacol. 8, 620–626

(2008).

(5) J.G. Elvin, R.G. Couston, and C.F. van der

Walle, Int. J. Pharm. 440, 83–98 (2013).

(6) D.M. Ecker, S.D. Jones, and H.L. Levine,

mAbs 7, 9–14 (2015).

(7) G. Walsh, Nat. Biotechnol. 32, 992–1000

(2014).

(8) G. Walsh, Nat. Biotechnol. 28, 917–924

(2010).

(9) A. Beck and J.M. Reichert, mAbs 5,

621–623 (2013).

(10) K. Sandra, I. Vandenheede, and P.

Sandra, J. Chromatogr. A 1335, 81–103

(2014).

(11) K. Sandra, I. Vandenheede, and P.

Sandra, LCGC Europe May Supplement,

10–16 (2013).

(12) A. Beck, S. Sanglier-Cianférani, and A.

Van Dorsselaer, Anal. Chem. 84, 4637–

4646 (2012).

(13) A. Beck, E. Wagner-Rousset, D. Ayoub,

A. Van Dorsselaer, and S. Sanglier-

Cianférani, Anal. Chem. 85, 715–736

(2013).

(14) E. Dumont, I. Vandenheede, P. Sandra,

K. Sandra, J. Martosella, P. Duong, M.

Joseph, Agilent Technologies Application

Note 5991-5124EN (2014).




Note 5991-5125EN (2014).




Note 5991-5135EN (2014).

(17) www.gene.com

(18) A. Staub, D. Guillarme, J. Schappler, J.L.

Veuthey, and S. Rudaz, J. Pharm. Biomed.

Anal. 55, 810–822 (2011).

(19) S. Fekete, J.L. Veuthey, and D. Guillarme,


(20) G. Chevreux, N. Tilly, and N. Bihoreau,

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


Belgium).

Pat Sandra is Chairman at the

Research Institute for Chromatography

(RIC, Kortrijk, Belgium) and Emeritus

Professor at Ghent University (Ghent,

Belgium).


Sandra et al.


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


www.palsystem.com


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,


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.


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



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).


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



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).


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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.

References(1) A. Beck, T. Wurch, C. Bailly, and N.

Corvaia, Nat. Rev. Immunol. 10, 345–352

(2010).

(2) A. Beck, J.F. Haeuw, T. Wurch, L.

Goetsch, C. Bailly, and N. Corvaia,

Discov. Med. 10, 329–339 (2010).

(3) A. Beck and J.M. Reichert, MAbs 6,

15–17 (2014).

(4) A. Beck, P. Senter, and R. Chari, MAbs 3,

331–337 (2011).

(5) A. Beck, J. Lambert, M. Sun, and K. Lin,

MAbs 4, 637–647 (2012).

(6) C. Klinguer-Hamour, P. Strop, D.K. Shah,

L. Ducry, A. Xu, and A. Beck, MAbs 6,

18–29 (2014).

(7) P.D. Senter and E.L. Sievers, Nat.

Biotechnol. 30, 631–637 (2012).

(8) M.M. Sun, K.S. Beam, C.G. Cerveny, K.J.

Hamblett, R.S. Blackmore, M.Y. Torgov,

F.G. Handley, N.C. Ihle, P.D. Senter, and

S.C. Alley, Bioconjug. Chem. 16, 1282–

1290 (2005).

(9) L.N. Le, J.M. Moore, J. Ouyang, X. Chen,

M.D. Nguyen, and W.J. Galush, Anal.

Chem. 84, 7479–7486 (2012).

(10) J.F. Valliere-Douglass, S.M. Hengel, and

L.Y.Pan, Mol. Pharm. 12, 1774–1783 (2015).

(11) L.Y. Pan, O. Salas-Solano, and J.F.

Valliere-Douglass, Anal. Chem. 86,

2657–2664 (2014).

(12) A. Beck, E. Wagner-Rousset, D. Ayoub,

A. Van Dorsselaer, and S. Sanglier-

Cianferan, Anal. Chem. 85, 715–736

(2013).

(13) A. Beck, MAbs 6(1), 30–33 (2014).

(14) A. Wakankar, Y. Chen, Y. Gokarn,

and F.S. Jacobson, MAbs 3, 161–172

(2011).

(15) J.F. Valliere-Douglass, W.A. McFee, and

O.Salas-Solano, Anal. Chem. 84,

2843–2849 (2012).

(16) S.M. Hengel, R. Sanderson, J. Valliere-

Douglass, N. Nicholas, C. Leiske, and

S.C. Alley, Anal. Chem. 86, 3420–3425

(2014).

(17) J. Chen, S. Yin, Y. Wu, and J. Ouyang,

Anal. Chem. 85, 1699–1704

(2013).

(18) A. Dyachenko, G. Wang, M. Belov, A.

Makarov, R.N. de Jong, E.T. van den

Bremer, P.W. Parren, and A.J. Heck,

Anal. Chem. 87, 6095–6102 (2015).

(19) F. Debaene, A. Boeuf, E. Wagner-

Rousset, O. Colas, D. Ayoub, N. Corvaia,

A. Van Dorsselaer, A. Beck, and S.

Cianferani, Anal. Chem. 86, 10674–10683

(2014).

(20) M.C. Janin-Bussat, M. Dillenbourg,

N. Corvaia, A. Beck, and C. Klinguer-

Hamour, J. Chromatogr. B 981–982, 9–13

(2015).

(21) E. Wagner-Rousset, M.C. Janin-Bussat,

O. Colas, M. Excoffier, D. Ayoub,

J.F. Haeuw, I. Rilatt, M. Perez, N. Corvaia,

and A. Beck, MAbs 6, 173–184 (2014).

(22) D. Lebert, G. Picard, C. Beau-Larvor,

L. Troncy, C. Lacheny, B. Maynadier,

W. Low, N. Mouz, V. Brun, C. Klinguer-

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.




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.



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


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.



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


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.



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.



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

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2

00 1 10 100 1000

5678

43210

0 1 10 100 1000

7

6

5

4

3

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00 1 10 100 1000

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00 1 10 100 1000

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10

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00 1 10 100 1000

12

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6

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00 1 10 100 1000

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00 1 10 100 1000

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Rela

tive a

bu

nd

an

ce (

Da)

Rela

tive a

bu

nd

an

ce (

Da)

Rela

tive a

bu

nd

an

ce (

Da)

Rela

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.



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.

References(1) J.J. Thomson, Proceedings of the Royal

Society of London Series a-Containing

Papers of a Mathematical and Physical

Character 89, 1–20 (1913).

(2) F.W. Aston, Nature 105, 547–547, (1920).

(3) K. Biemann, Proc. Natl. Acad. Sci. U.S.A.

104, 10310–10313 (2007).

(4) R. Aebersold and M. Mann, Nature 422,

198–207 (2003).

(5) O. Fiehn, Plant Mol. Biol. 48, 155–171 (2002).

(6) C.K. Meng, M. Mann, and J.B. Fenn, Z.

Phys. 10, 361–368 (1988).

(7) M. Karas and F. Hillenkamp, Anal. Chem.

60, 2299–2301 (1988).

(8) J.K. Eng, A.L. McCormack, and J.R.I.

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).

(11) J.A. Loo, H.R. Udseth, and R.D. Smith,

Biomed. Environ. Mass Spectrom. 17,

411–414 (1988).

(12) M. Przybylski and M.O. Glocker, Angew.

Chem. Int. Ed. 35, 806–826 (1996).

(13) J.A. Loo, Int. J. Mass spectrom. 200,

175–186 (2000).

(14) M.C. Fitzgerald, I. Chernushevich, K. G.

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).

(18) A.B. Kanu, P. Dwivedi, M. Tam, L. Matz, and

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).

(20) V. Katta and B.T. Chait, J. Amer. Chem.

Soc. 115, 6317–6321 (1993).

(21) K.D. Rand, M. Zehl, and T.J.D. Jorgensen,

Acc. Chem. Res. 47, 3018–3027 (2014).

(22) D. Houde, S.A. Berkowitz, and J.R. Engen,

J. Pharm. Sci. 100, 2071–2086 (2011).

(23) A. Sinz, Mass Spectrom. Rev. 25, 663–

682 (2006).

(24) A. Sinz, J. Mass Spectrom. 38, 1225–1237

(2003).

(25) A. Leitner, T. Walzthoeni, and R. Aebersold,

Nature Protocols 9, 120–137 (2014).

(26) C. Bich, S. Maedler, K. Chiesa, F.

DeGiacomo, N. Bogliotti, and R. Zenobi,

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.


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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.



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).


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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.


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


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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.

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[rhTRAIL]=10 ng/mL [rhTRAIL]=10 ng/mL

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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.


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


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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.

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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.


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


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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.


Advances in Biopharmaceutical Analysis – OCTOBER 2015 43

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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.


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).

(2) G. Hopfgartner, A. Lesur, and E.

Varesio, Trends Anal. Chem. 48, 52–61

(2013).

(3) I. van den Broek, W.M. Niessen, and

W.D. van Dongen, J. Chromatogr. B

929, 161–179 (2013).

(4) K.J. Bronsema, R. Bischoff, and N.C.

van de Merbel, J. Chromatogr. B 893-

894, 1–14 (2012).

(5) D. Wilffert, R. Bischoff, and N.C. van de

Merbel, Bioanalysis 7, 763–779 (2015).

(6) K.J. Bronsema, R. Bischoff, and N.C.

van de Merbel, Anal. Chem. 85, 9528–

9535 (2013).

(7) K.J. Bronsema, R. Bischoff, W.W.M.P.

Pijnappel, A.T. van der Ploeg, and

N.C. van de Merbel, Anal. Chem. 87,

4394–4401 (2015).

(8) D. Wilffert, C.R. Reis, J. Hermans, N.

Govorukhina, T. Tomar, S. de Jong,

W.J. Quax, N.C. van de Merbel, and R.

Bischoff, Anal. Chem. 85, 10754–10760

(2013).

(9) V. Kumar, D.R. Barnidge, L.S. Chen,

J.M. Twentyman, K.W. Cradic, S.K.

Grebe, and R.J. Singh, Clin. Chem. 56,

306–313 (2010).

(10) J.R. Whiteaker, L. Zhao, H.Y. Zhang,

L.C. Feng, B.D. Piening, L. Anderson,

and A.G. Paulovich, Anal. Biochem.

362, 44–54 (2007).

(11) K.J. Bronsema, R. Bischoff, M.P.

Bouche, K. Mortier, and N.C. van de

Merbel, Bioanalysis 7, 53–64 (2015).

(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.


van de Merbel

ES683327_LCESUPP1015_044.pgs 10.01.2015 20:53 ADV blackyellowmagenta


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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.



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).



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.



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.

References

(1) F. Wang, D. Richardson, and M.

Shameem, BioPharm Int. 28, 32–38

(2015).

(2) Q. Zhang, A.M. Goetze, H. Cui, J. Wylie,

S. Trimble, A. Hewig, and G.C. Flynn,

mAbs 6, 659–670 (2014).

(3) C.E. Doneanu, A. Xenopoulos, K. Fadgen,

J. Murphy, S.J. Skilton, H. Prentice, M.

Stapels, and W. Chen, mAbs 4, 24–44

(2012).

(4) M. Pavlovic, E. Girardin, L. Kapetanovic,

K. Ho, and J.H. Trouvin, Horm. Res. 69,

14–21 (2008).

(5) J. Geigert, The Challenge of

CMC Regulatory Compliance for

Biopharmaceuticals and other Biologics

(Springer Science & Business Media,

Heidelberg, Germany, 2013).

(6) S.X. Gao, Y. Zhang, K. Stansberry‑

Perkins, A. Buko, S. Bai, V. Nguyen, and

M.L. Brader, Biotechnol. Bioeng. 108,

977–982 (2011).

(7) M.R. Schenauer, G.C. Flynn, and A.M.

Goetze, Anal. Biochem. 428, 150–157

(2012).

(8) J.H. Thompson, W.K. Chung, M. Zhu, L.

Tie, Y. Lu, N. Aboulaich, R. Strouse, and

W. Mo, Rapid Comm. Mass Spectrom. 28,

855–860 (2014).

(9) V. Reisinger, H. Toll, R.E. Mayer, J. Visser,

and F. Wolschin, Anal. Biochem. 463, 1–6

(2014).

(10) C.E. Doneanu and W. Chen, Methods

Mol. Biol. 1129, 341–350 (2014).

(11) K. Bomans, A. Lang, V. Roedl, L. Adolf,

K. Kyriosoglou, K. Diepold, G. Eberl, M.

Mølhøj, U. Strauss, C. Schmalz, R. Vogel,

D. Reusch, H. Wegele, M. Wiedmann, and

P. Bulau, PLoS One 8, e81639 (2013).

(12) M.R. Schenauer, G.C. Flynn, and A.M.

Goetze, Biotechnol. Bioeng. 29, 951–957

(2013).

(13) K. Sandra, M. Moshir, F. D’hondt, R.

Tuytten, K. Verleysen, K. Kas, I. François,

and P. Sandra, J. Chromatogr. B 877,

1019–1039 (2009).

(14) M.P. Washburn, D.A. Wolters, and J.R.

Yates, Nat. Biotechnol. 19, 242–247

(2001).

(15) D.A. Wolters, M.P. Washburn, and J.R.

Yates, Anal. Chem. 73, 5683–5690

(2001).

(16) N. Delmotte, M. Lasaosa, A. Tholey, E.

Heinzle, and C.G. Huber, J. Proteome

Res. 6, 4363–4373 (2007).

(17) M. Gilar, P. Olivova, A.E. Daly, and J.C.

Gebler, Anal. Chem. 77, 6426–6434

(2005).

(18) G. Vanhoenacker, I. Vandenheede, F.

David, P. Sandra, and K. Sandra, Anal.

Bioanal. Chem. 407, 355–366 (2015).

(19) K. Sandra, K. Mortier, L. Jorge, L.C.

Perez, P. Sandra, S. Priem, S. Poelmans,

and M.P. Bouche, Bioanalysis 6, 1201–

1213 (2014).

Koen Sandra is Director

at the Research Institute for

Chromatography (RIC, Kortrijk,

Belgium).

Alexia Ortiz is a Proteomics

Researcher at the Research Institute


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.


Advances in Biopharmaceutical Analysis – OCTOBER 2015 49


When it comes to addressing biosimilar and monoclonal

antibodies (mAb) quantitation in biological matrices, research

groups and pharmaceutical organizations face two major

challenges: poorly developed pretreatment processes in terms of

selectivity and repeatability, and a compromise on speed and/or

sensitivity of liquid chromatography tandem mass spectrometry

(LC–MS–MS) systems. One could consider the enzyme-linked

immunosorbent assay (ELISA) technology as an alternative

solution, but lack of development time, increased costs, high

failure rates, and cross reactivity compels scientists to look at

mass spectrometry-based solutions.

Shimadzu L i fe Science Research Center has worked

relentlessly on the establishment of a universal bioanalytical

pretreatment method for IgG derived mAbs that is easy and

more selective. The current proteolysis methods (proteolysis

is the breakdown of proteins into smaller polypeptides) can

make identification of signature peptides among a gamut of

peptides very dif ficult, thereby decreasing the quantitative

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.


50 Advances in Biopharmaceutical Analysis – OCTOBER 2015


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.


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