MS-based conformation analysis of recombinant proteins in design, optimization and development of biopharmaceuticals

Accepted Manuscript MS-based conformation analysis of recombinant proteins in design, optimization and development of biopharmaceuticals Devrishi Goswami, Jun Zhang, Pavel V. Bondarenko, Zhongqi Zhang PII: DOI: Reference:

S1046-2023(17)30454-1 https://doi.org/10.1016/j.ymeth.2018.04.011 YMETH 4448

To appear in:

Methods

Received Date: Revised Date: Accepted Date:

6 February 2018 10 April 2018 12 April 2018

Please cite this article as: D. Goswami, J. Zhang, P.V. Bondarenko, Z. Zhang, MS-based conformation analysis of recombinant proteins in design, optimization and development of biopharmaceuticals, Methods (2018), doi: https:// doi.org/10.1016/j.ymeth.2018.04.011

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MS-based conformation analysis of recombinant proteins in design, optimization and development of biopharmaceuticals Devrishi Goswami, Jun Zhang, Pavel V. Bondarenko* and Zhongqi Zhang* Process Development, Amgen, 1 Amgen Center Drive, Thousand Oaks, CA 91320

*Corresponding author Zhongqi Zhang, [email protected] Pavel V. Bondarenko, [email protected]

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1. Introduction 2. Method overview 2.1 Native MS 2.2 HDX-MS 2.3 Covalent labeling 2.4 Cross-linking 2.5 Limited proteolysis 3. Application in developing protein therapeutics 3.1 Drug discovery, design and optimization 3.1.1 Antibody-antigen interaction 3.1.2 Epitope/paratope mapping 3.1.3 Impact of protein conjugation 3.1.4 Understanding heterogeneity 3.1.5 Molecular engineering 3.2 Cell Line, Cell Culture and Purification Development 3.2.1 Impact of quality attributes 3.2.2 Comparability/biosimilarity study 3.3 Formulation development 3.3.1 Conformational stability 3.3.2 Dimer, oligomer and aggregate formation 3.3.3 Viscosity 3.3.4 Protein-excipient interactions 4. Conclusion and future outlook

Abstract Mass spectrometry (MS)-based methods for analyzing protein higher order structures have gained increasing application in the field of biopharmaceutical development. The predominant methods used in this area include native MS, hydrogen deuterium exchange-MS, covalent labeling, cross-linking and limited proteolysis. These MS-based methods will be briefly described in this article, followed by a discussion on how these methods contribute at different stages of discovery and development of protein therapeutics.

Keywords: biopharmaceutical; antibody; higher order structure; native mass spectrometry; hydrogen/deuterium exchange; oxidative footprinting

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1. Introduction During the last decade, recombinant therapeutic proteins occupy a major portion of the approved drugs globally. They are expected to grow not only by numbers but also in the overall market share. Biotechnology product sales is expected to grow from 25% in 2016 to 30% ($326 Billion worldwide sales) in 2022 and 52% of the top 100 products by value will be biologics (EvaluatePharma: World Preview 2017, Outlook to 2022, 10th ed. (2017)). All major pharmaceutical companies are devoting more of their pipeline assets towards biologics. With the advancement of recombinant DNA/hybridoma technology and XenoMouse method [1], large quantity of biologics is produced routinely for clinical and commercial use. The rise in demand for biologic drug is partly due to its superior specificity, high efficacy, long life in circulation and fewer side effects. However, the efficacy, clearance and immunogenicity of biologics are highly dependent on attributes such as amino acid sequence, post translational modifications, process and storage induced modifications, and higher order structure (HOS). Unlike synthetic small-molecule drugs, therapeutic proteins are produced recombinantly inside a living organism involving a highly complex bioprocessing machinery. Each step of production has a dramatic impact on the downstream protein quality, which in turn influences the efficacy, clearance and safety of the biological product. The production, purification and formulation processes are typically optimized to minimize the exposure of harsh conditions such as extreme pH, elevated temperature, high detergent concentration, as well as mechanical and other stresses. If there are deviations from the optimized process, however, the exposure to unusually harsh conditions can lead to covalent (chemical) and non-covalent modifications causing alterations of HOS. Most changes in covalent structure involve a change in mass, which can be conveniently detected by mass spectrometry (MS). Because of its superior sensitivity, selectivity, resolution and robustness, MS has evolved as an essential tool for product characterization from early discovery phase to clinical and commercial release stage [2]. Many product quality characteristics (e.g., aggregation or chemical degradation) can be either optimized or exacerbated during each stage of the recombinant protein manufacturing. In addition to conventional chromatographic techniques, MS evaluation of protein quality at each stage is used to identify specific processing steps that lead to increased levels of sequence variants, post-translationally modified forms, degradation products, and impurities, etc. Additionally, MS is routinely used during formulation development to identify formulation matrices that provide optimal protection from chemical degradation and aggregation, and hence protect from activity loss associated with it. It is also used to monitor chemical modifications such as methionine oxidation, asparagine

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deamidation, and aspartate isomerization following storage of formulated bulk solutions or lyophilized products under moderate and accelerated conditions. Since early 1990’s, besides monitoring covalent structural changes, several MS-based techniques have been developed to monitor secondary, tertiary, or quaternary structural features, namely HOSs, of proteins [3]. First, protein quaternary structure and protein-protein interaction can be probed by MS analysis in a native-like condition, in which non-covalent protein-protein interactions are preserved. In addition, protein unfolding, change in HOS or protein-protein interaction may lead to alteration in charge distribution and collision cross-section, detectable by modern mass spectrometric techniques. Secondly, because noncovalent structural changes do not involve a mass difference, a covalent reaction (or labeling) step may be applied to the protein before MS analysis. The reaction/labeling step creates mass changes that are detectable by mass spectrometry, while the information contained in these mass changes yields indirect HOS information. These techniques, including covalent modification, limited proteolysis, crosslinking, and especially hydrogen/deuterium exchange (HDX), are getting increasingly more applications in the development of therapeutic proteins. This review will focus on HOS analysis of protein therapeutics by MS. With a brief introduction to the methods and instrumentations, we will discuss how HOS analysis via MS contributes at different stages of drug discovery and development using specific examples. Finally, we will discuss the future trend and criticality of HOS analysis by MS in biopharmaceuticals development.

2. Method Overview

2.1 Native MS During electrospray ionization (ESI), if the ionization condition is soft enough, the weak noncovalent interactions that hold together the overall folded structure of a protein or a protein complex can often be preserved into the gas phase, and therefore be analyzed by mass spectrometry [4]. This soft ionization requires the protein solution to be ionized from a “native” state. This is possible by using an aqueous solution of volatile buffer at a neutral pH (most commonly ammonium acetate, but also ammonium bicarbonate or triethylammonium bicarbonate). As the first step, the protein must be exchanged into the appropriate volatile buffer before native MS analysis. Due to the folded nature of the gas-phase protein molecules, native MS generates fewer charges comparing to denaturing conditions. The fact that folded state generates fewer charges can also be conveniently used to evaluate the compactness of the protein folding. 4

Native MS can also be connected downstream to size-exclusion [5-7] or ion-exchange chromatography [8, 9], if MS-compatible buffer is used in the mobile phase. This is usually achieved by using a volatile buffer in the mobile phase, and in the case of ion-exchange, using a pH gradient instead of a salt gradient for elution. Native MS, combined with charge-state distribution or ion-mobility (see below) measurement, is able to evaluate conformations of protein isoforms separated by size-exclusion and ionexchange chromatography. Most native MS experiments are performed using one of the following three types of configurations. Time-of-flight (TOF) and quadrupole time-of-flight (Q-TOF) analyzer: A TOF MS [10] is traditionally the most preferred analyzer for native MS study. In a TOF analyzer, m/z is determined by measuring the time the ions take to reach the detector traveling through a field-free flight tube under high vacuum. In a slightly modified version, Q-TOF, a mass-selecting quadrupole (Q) analyzer and a TOF analyzer are separated by a collision cell. This setup allows tandem-MS (MS/MS) experiments, in which a precursor ion is selected in the quadrupole, dissociated in the collision cell via collision-induced dissociation (CID), and the product ions are analyzed in the TOF. Ion Mobility-MS (IM-MS): Ion mobility (IM) [11] is a technique that allows the separation of the ions based on their charge as well as gas-phase collision cross-section (CCC). The separation occurs in a gas-filled cell located between the quadrupole and the TOF. Ions with different sizes, crossing the cell under the influence of an electric field, experience different interactions with the gas molecules, thus having different velocities. Ions with larger or extended conformations have larger CCCs, and therefore drift more slowly than those with more compact shapes. Ion-mobility, combined with a Q-TOF analyzer, not only measures the mass of the protein molecules, but also separates the molecules based on their gasphase shapes, therefore opening the door to a new dimension of HOS information. Orbitrap and other mass spectrometers: The Orbitrap™ is a special type of mass analyzer, consisting of an external barrel-shaped electrode and an internal spindle-shaped one [12]. An electrostatic voltage, applied to the central electrode, generates a field that forces the ions to oscillate along the central electrode with a frequency determined by the m/z ratio. A Fourier transform converts the current generated by the oscillating ions into frequencies and intensities yielding the mass spectrum. In 2012, Rose et al. [13] described for the first time the use of a modified Orbitrap mass analyzer (EMR- Extended mass range) to measure non-covalent protein assemblies of molecular weights approaching one megadalton using native MS, with a sensitivity allowing the detection of single ions. Recently, Exactive PlusTM (Thermo Fisher Scientific, Bremen, Germany) Orbitrap instrument has been available for usage in native MS mode. The instrument is equipped with a higher-energy collisional dissociation (HCD) cell in which the ions are stored for improved transmission and desolvation. The main advantage of this type of

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instrument is the ultra-high mass resolution. Other less frequently used mass analyzers include Fouriertransform ion cyclotron resonance (FTICR) [14, 15]. Analyzing mass spectra of intact proteins usually involves deconvolution of the spectrum to a zerocharged mass domain spectrum. These software tools are usually provided by the instrument manufacturer. Analysis of ion-mobility data is not discussed here due to its complexity; the topic has been discussed extensively in several review papers [16, 17].

2.2

HDX-MS

When a protein (or protein complex) is exposed to a deuterated solution, backbone amide hydrogen atoms on the protein will exchange with deuterium atoms in the solution. Backbone amide hydrogen atoms on the surface of the protein that are not involved in hydrogen bonding, exchange with solution deuterium atoms faster than those in the interior of the protein or involved in strong hydrogen bonding. As a result, monitoring HDX behavior will yield information on protein conformation and dynamics [18-20]. Exchange of a hydrogen atom with a deuterium atom yields a mass change, which can be conveniently determined by mass spectrometry. Before MS is used for HDX in early 1990’s [21], hydrogen exchange measurements were mostly performed by NMR [20], which provides structural information at singleresidue resolution. However, application of NMR is limited to small proteins. In addition, large sample requirement and lengthy resonance assignment process also limits the wide applicability of NMR. In recent years, HDX-MS has become the method of choice for analyzing conformational changes in protein molecules due to its sensitivity, spatial resolution, automation, and applicability to large proteins. In a typical “bottom-up” HDX-MS workflow [21], a protein sample is incubated in a deuterium-containing buffer for certain time periods, and then quenched by bringing the reaction to low pH and temperature (pH 2.5, 0-4°C). The deuterium labeled sample is then digested by an acid protease, followed by chromatographic separation and mass spectrometric analysis. Depending on the questions to be answered, two types of labeling experiments, including continuous labeling and pulsed labeling, are often applied [22]. In a continuous labeling experiment, the protein sample is incubated in a deuterated buffer for certain time periods to allow deuterium labeling. Because the deuterium labeling reaction is accumulative during the labeling time period, the deuterium uptake reflects the entire history of the protein solvent accessibility and conformation/dynamics within that time period. Continuous labeling is usually used to study protein conformation/dynamics under the equilibrated state. In a pulsed labeling experiment, on the other hand, protein conformation is first perturbed in some manner, such as by exposing to a chemical denaturant, for certain periods of time, and then exposed to deuterated buffer for a very brief time period (a labeling pulse, typically 10 s or less) [22, 6

23]. During this short labeling time, the conformational changes are negligible compared to those induced by the perturbation. As a result, the deuterium uptakes in a pulsed labeling experiment represent snapshots of the protein conformation upon perturbation in a time-resolved manner. Pulsed labeling is used to follow the kinetics of a conformational change. The deuterium labeled protein after quenching can be analyzed directly by MS to determine the exchange profile of the intact protein. Furthermore, the labeled protein can be analyzed using “bottomup” approach, by which the protein is digested into small pieces to localize the deuterium incorporation at the peptide level. While the assignment of deuterium to individual residues from MS/MS data is typically difficult due to the deuterium scrambling during the collisional activation in gas phase, recent reports have indicated that the low energy activation methods, e.g., electron-capture dissociation (ECD) and electron-transfer dissociation (ETD), are promising with minimal deuterium scrambling, so that ECD and ETD can be implemented in an MS method to monitor the conformational dynamics of proteins at singleresidue resolution [24, 25] [26] [27]. On top of that, ECD and ETD can be applied to intact protein directly to enable “top-down” HDX-MS [28]. Top-down HDX-MS has gained popularity because of its advantage of conformer-specific conformational analysis, no requirement of proteolytic digestion, and dramatically reduced level of back exchange. However, challenges remain for larger proteins with multiple disulfide bonds, which are common in therapeutic proteins including mAbs [29, 30], as well as potential deuterium scrambling during ionization/fragmentation. HDX experiments can be performed in a variety of ways. Traditional HDX reactions are carried out at seconds to hours labeling time scales. Several groups reported both top-down and bottom-up HDX-MS workflow at millisecond time scale to monitor the weakly structured dynamic regions and sub second conformational transitions [25, 28, 31-35]. To address the conformational behavior and aggregation propensity of protein molecule due to freezing, Zhang et al. developed a HDX-MS workflow in a frozen state [36, 37]. HDX-MS analysis of protein sample at high concentration was addressed by lyophilization of protein sample and subsequent reconstitution in deuterated labeling buffer [38] or a dilution-free dialysis based method [39]. In order to better understand protein conformational stability in amorphous solid state and lyophilization-induced protein degradation, a solid state HDX-MS work flow has been developed and extensively used by Elizabeth Topp and coworkers for studying conformational stabilities of lyophilized protein products [40-44]. A major advancement of the HDX-MS technology that enables its wide application is the development of the automation platform. With many repetitive steps, an HDX-MS experiment is ideally automated. The most popular platform for real time HDX-MS automation is the HTS Twin PAL autosampler (CTC Analytics, Zwingen, Switzerland, supplied in USA by Leap Technologies which is recently acquired by Trajan Scientific and Medical, Victoria, Australia). Several software packages are 7

available in the market to control the automated operation, namely, Cycle Composer (CTC Analytics, Zwingen, Switzerland) and its newer developments - HDX Director and Chronos (LEAP technologies) etc. [45-50]. Another major advancement of HDX-MS technology involves automated data processing. Bottom up HDX-MS experiments typically generate hundreds of overlapping peptides from various time points over a range of labeling periods. As part of HDX-MS workflow, data processing involves analyzing a large number of mass spectra for deuterium labeling information [51]. This is an extremely time consuming and labor-intensive task if performed manually. Over the years, many data processing software suites were developed with faster processing times and better accuracy, namely, AutoHD [52], HDExaminer (Sierra Analytics)[46, 53], HX-Express [54], DEX [55], Hydra [56], HeXicon [57], EX-MS [58], HDXworkbench/desktop [59-61], DynamX (Waters), MassAnalyzer [50], HDX Finder [62], HDsite [63], Mass Spec studio [64] and others [65]. Most of these software tools calculate deuterium uptakes automatically based on centroid mass of the labeled peptide determined from the mass spectra. By Monte Carlo simulation using a comprehensive HDX model, MassAnalyzer (available from Thermo Scientific in Biopharma Finder) developed by Zhang et al. is able to determine possible protection factor of each backbone amide hydrogen in a fully automated fashion [50]. Deuterium uptakes determined from the peptide centroid mass, however, do not reflect all the information contained in the mass spectrum of a deuterated peptide. Protein conformation and dynamic information is also contained within the isotope distribution pattern [51], which is often neglected. For example, Zhang et al. [66], using HDX study of Ca2+ binding to an N-terminus truncated DREAM (downstream regulatory element antagonist modulator), reported that simply calculating the centroid mass of target peptides can lead to a wrong conclusion. Therefore, besides centroid calculation, it is also beneficial to examine the isotopic distributions in the mass spectra of the deuterated peptides/proteins.

2.3 Covalent labeling Complementary to HDX, amino acid side chains can be labeled by a chemical reagent when they are exposed to solvent [67]. There are two general types of covalent labeling reagents - amino acid specific and non-specific. For example, acetic anhydride can be used to label solvent accessible lysine residues, GEE (glycine ethyl ester) to label Asp and Glu residues [68], NEM (N- Ethylmaleimide) to label Cys residues, etc [67]. Amino acid non-specific approaches include oxidative labeling by hydroxyl radicals [69]. Covalent labeling approach for conformational analysis offers advantage that the labeling are irreversible. Therefore, the protein can be digested with any protease of choice and analyzed by any LC and MS method of choice. As a result, the labeling can be more extensively characterized. Unlike HDX, 8

however, extensive labeling will change the protein conformation and produce incorrect conclusion if care is not taken. Therefore, the labeling reaction must either be controlled so that the extent of covalent labeling is minimal, or the labeling reaction must complete within a very short period of time before any conformational change can occur. Of particular interest for therapeutic protein conformational analysis is oxidative labeling by hydroxyl radicals. Oxidative labeling is a powerful technique to map the interface between domains and other folded structural elements of macromolecules and to map the conformational changes due to proteinprotein or protein-ligand interactions. The method was developed by Chance and co-workers [70-72] who exposed protein solutions to high-energy radiation produced in a synchrotron to ionize water and form short-lived hydroxyl radicals (·OH) by H+ loss. Hydroxyl radicals can react with side-chain sites in proteins that are solvent accessible; the resultant mass shift marks the site of modification and the modification level can be used to track the solvent accessibility of that site. The irreversibly modified protein products can be detected and quantified by liquid chromatography/tandem MS (LC-MS/MS), typically after proteolysis, providing measures of surface accessibility for specific side chains in the protein structure. Hydroxyl radical can be generated by several methods, such as chemical methods [73], radiolysis of water [74, 75] or photolysis of hydrogen peroxide [76]. A modified version of oxidative footprinting, named fast photochemical oxidation of proteins (FPOP), was developed by two groups at a similar time [77, 78]. In this method, an excimer laser is used to photolyze hydrogen peroxide, which is added in low concentration to the protein solution, to form hydroxyl radicals. The use of a pulsed laser and a radical scavenger limits radical lifetimes, ensuring labeling on a microsecond time scale, faster than most protein unfolding [79]. Gross lab further developed the labeling approach and reported using iodine radical (photolysis of iodobenzoic acid at 248 nm) to selectively modify only histidine and tyrosine residues [80], or laser-initiated radical trifluoromethylation (·CF3), which can modify broad range of residues including Gly, Ala, Ser, Thr, Asp, and Glu, which are relatively silent with regard to ·OH [81]. The variety of oxidation reactions mediated by hydroxyl radicals complicate MS data, and therefore computing different level of oxidation for each peptide is immensely time consuming. To speed up this process, ProtMapMS software [82] was developed for automated identification and quantification of the extent of modifications. ProtMapMS first identifies both modified and unmodified forms of the peptide and then extracts their corresponding chromatograms for the target peptide precursor ions based on the window defined by a user around the mass value of interest (in ppm). Finally, it returns a dose response curve and oxidation rate of peptides.

2.4 Cross-linking 9

Cross-linking associated with mass spectrometry (XL-MS) is a powerful technique for protein structure elucidation. By using bifunctional cross-linkers with different lengths, one can determine the maximum distance between two residues and therefore obtain 3-dimensional information of the molecule. This technique can be used to characterize protein complex, or determine a low-resolution 3-dimensional structure when combined with molecular dynamic simulation [83]. XL-MS provides several layers of information, for example, identifying protein–protein contacts confirms physical proximity between subunits, localizing the side chains that are cross-linked restricts this proximity to certain regions (e.g., domains or even single helices or loops), and finally, the structure of the cross-linked side chains and the cross-linker moiety impart a distance restraint that can be used for molecular modeling purposes [84]. Typically, a purified protein or protein-protein complex (e.g., Ag-Ab) is incubated with a crosslinking reagent to form covalent bonds between reactive surface-exposed amino acid side chains. After proteolytic digestion, the resulting peptides are analyzed by LC-MS/MS. Computational analysis of the MS/MS data enables sequence assignment of the cross-linked peptides as well as the localization of the exact cross-linking sites. Commonly used cross-linking reagents are homobifunctional active esters such as disuccinimidyl suberate (DSS) or bis(sulfosuccinimidyl) suberate (BS3) that induce nucleophilic attacks on primary amines and thus rely on the coupling of lysine residues. Another cross-linking reagent, homobifunctional dihydrazides, relies on cross-linking chemistry specific for carboxyl groups [85].

2.5 Limited Proteolysis Similar to covalent labeling, the protein can be exposed to a protease at a native condition. The residues on the surface of the protein have access to the protease and therefore nearby backbone can be hydrolyzed. Residues in the interior of the molecule, however, are protected from the protease. The site of the cleavage can be determined by mass spectrometry to yield conformational information [86]. Also similar to covalent labeling, the extent of limited proteolysis must be well controlled to avoid incorrect conclusion due to change of conformation after the cleavage. At the end of 1990s – beginning 2000s, limited proteolysis in combination with mass spectrometry have been actively applied to probe conformational features and stability of proteins. The technique provided important information on the structure and dynamics, complementing the results of biophysical and spectroscopic techniques. Modified, mutated, truncated protein variants released the proteolysis products at different rate and sometimes cleaved at different sites as compared to the wildtype protein, providing useful information about conformation differences. Several enzymes with different specificities were typically required to achieve useful coverage. For example, surface topology 10

of synthetic fibrils obtained from intact and truncated beta2-microglobulin was investigated using trypsin, chymotrypsin, and endoprotease Asp-N followed by LC-MS. Enzymatic digestions were all performed in 100 mM sodium phosphate (pH 7.5) at 37°C using enzyme-to-substrate ratios ranging between 1:100 and 1:1000 (w/w) [87]. Tertiary structure of the Minibody, composed of heavy chain domains of mouse immunoglobulin, was studied by limited proteolysis with trypsin, chymotrypsin, elastase, subtilisin and endoproteinase Glu-C accompanied by LC-MS. The limited proteolysis was performed using pH 7.5, 37°C, enzyme-to-substrate ratio of 1:2000, and the enzymatic hydrolyses were monitored on a timecourse basis [88]. The methodology was applied to check the folding state of a protein and characterize mutational effects on protein conformation and stability [89]. Even with multiple proteases, the technique could provide only limited structural resolution and was eventually replaced by higher resolution techniques described in this review. One advantage of the technique is that it requires small amounts (micrograms) to perform experiments.

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Applications in Developing Protein Therapeutics

Bringing a protein drug from laboratory bench to commercial manufacturing is a long and complex process. Figure 1 shows a simplified process of developing a protein drug. First, during drug discovery, design and optimization, several potential protein drug candidates are designed to bind to the target, followed by selecting and optimizing the best candidate for clinical trials. After a candidate is selected, a cell line must be developed to stably express the recombinant protein therapeutics candidate. After that, upstream cell culture process and downstream purification process must be created and optimized to produce the recombinant protein as the drug substance. The drug substance needs to be formulated to ensure the stability over the shelf life and through the fill-and-finish process to manufacture the drug product. MS plays an important role in virtually every step of the process in analyzing covalent structural attributes. In terms of noncovalent structures, increasing number of MS-based methods have been used over the past 20 years due to their high sensitivity, fast turn-around-time, and spatial resolution compared to conventional techniques. Currently, these methods have been used to support therapeutic protein discovery and development. The involved activities include epitope and paratope mapping, impact of attributes on protein conformation, molecular heterogeneity, protein engineering, understanding aggregation/viscosity, protein-protein/protein-excipient interactions, and so on. In the following sections, progresses in each of these areas will be reviewed. Table 1 summarizes examples of conformation analyses of protein therapeutics during different stages of drug development.

3.1 Drug Discovery, Design and Optimization 11

During drug discovery, design and optimization, several potential protein drug candidates are designed to bind to the target, followed by selecting and optimizing the best candidate for clinical trials. To ensure efficacy, safety and developability of the candidate, this process requires the understanding, at the molecular level, of drug-target interaction, conformational stability, etc., in order to engineer the best possible molecule.

3.1.1

Antibody-antigen interaction

In recent years, antibodies and engineered immunoglobulin-like molecules have become the major types of protein biologics due to their many favorable properties including high specificity, superior stability, long half-lives, low toxicity, and easiness to make bispecific and drug conjugate, etc. These molecules bind to their target (antigen) strongly with high specificity. MS-based methods are playing increasingly important role in studying the interaction between antibodies and antigens. When a protein complex is analyzed by electrospray MS under a native-like condition, the complex can often be conserved into the gas phase. Carol Robinson’s group first reported the MS analysis of intact antigen-antibody complex in native condition using ammonium acetate as solvent to preserve the noncovalent interactions [90]. Using a combination of nanoflow electrospray and Q-TOF mass analyzer, they characterized the oligomeric nature of V antigen (from Yersinia pestis) in solution and the stoichiometry of the antigen-antibody (Ag-Ab) complex. Moreover, carefully increasing the collision energy in a stepwise fashion, they followed the collision-induced dissociation of antigen-antibody complex and further confirmed the binding stoichiometry and specificity of the interaction. The importance of extended m/z range made available by a TOF analyzer with collisional cooling to resolve such high macromolecular complex was underlined. They further characterized the influence of regional and segmental flexibility of mAb on antigen binding and stoichiometry [91]. Using MS in denaturing condition they deciphered the effect of antibody affinity and antigen variance on imunocomplex formation and showed that the stoichiometry of the antigen–antibody interaction is altered by restricting the movement of the Fc region [92]. Taking similar approach of ESI-TOF at a non-denaturing condition an Amgen group has characterized the in vitro stoichiometry of RANK ligand and denosumab, a fully humanized human IgG2 mAb. The study uncovered a stable complex of 590 kDa including two RANK ligand trimers cross-linked by three antibodies in a propeller configuration (Figure 2) [93]. In another study [94], Atmanene et al. used native MS including automated chip based nanoESI-MS and travelling wave ion mobility mass spectrometry (TWIMS) to characterize the formation of immunocomplexes involving murine mAb 6F4 and its humanized version raised against recombinant 12

human JAM-A (Junctional Adhesion Molecule-A) extra cellular domain. They used the TWIMS feature of Synapt HDMS (Waters, Manchester, UK) as an initial quality control step to analyze recombinant antigen batches (SF- purified from soluble fraction; IB- purified from insoluble fraction) in order to verify their structural homogeneity and to confirm that the folded conformation is maintained in the experimental conditions used for Ag-Ab binding assays. NanoESI-TWIMS analysis in denaturing conditions reveals that multiply charged ions of JAM-A IB display lower m/z with longer drift times than those detected for JAM-A SF. This observation confirms that JAM-A IB contains less disulfide bonds resulting in more extended protein conformations conferring lower mobilities (i.e., longer drift times) to protein ions. Taking together TWIMS analysis and other structural MS studies, this report concluded that in spite of having heterogeneous disulfide bridge pairings of recombinant JAM-A neither its native structure nor mAbs 6F4 recognition properties are altered. They stressed that due to low sample consumption, assay speed and automation, native MS played a strong role in Ag-Ab structure analysis, mAb drug development and lead optimization. Debaene et al. [95] also reported, using native MS and TWIMS, rapid assessment of bispecific antibody (bsAb) heterogeneity and bsAb-antigen complex characterization. IgG4 undergoes Fab-arm exchange (FAE) in which half molecules of two wild type IgG4 mAbs recombine to form heterotetramer. Leveraging the ion mobility feature, the group was able to monitor in real time FAE and subsequent bispecific mAbs formation in Hz6F4-2v3, a humanized IgG4. Moreover, native MS analysis of bsAb/JAM-A immune complexes revealed that bsAb can bind up to two antigen molecules, confirming that Hz6F4-2 preferentially binds dimeric JAM-A. In a recent tandem native MS work reported by Heck and coworker [96], submillion Da antibodyantigen complex has been characterized using an Orbitrap EMR (extended mass range) equipped with a high mass quadrupole mass selector and careful instrument optimization. Using this new tandem MS workflow, they can unambiguously probe the stoichiometry of antigen binding in such a large protein assembly.

3.1.2

Epitope/paratope mapping

Native MS can conveniently determine the stoichiometry of the antibody-antigen complex. To determine the binding interface (epitope or paratope mapping), however, a technique with spatial resolution is required. Epitope/paratope mapping is the identification and characterization of residues/motifs in the antigen (epitope) that are recognized and bound by its corresponding Fab arm of antibody (paratope). This work is important in the understanding of drug-target interaction to facilitate better design and optimization of 13

the drug. Traditionally this work has been performed on X-ray crystallography, in which the Fab domain of the antibody is co-crystallized with the antigen, and the binding interface was determined from the 3-D structure of the complex. Scanning mutagenesis, followed by a binding assay, is also used to determine the epitope. Both methods, however, are time-consuming and labor-intensive. MS-based methods are playing increasingly important role in epitope/paratope mapping. Pimenova et al. [97] reported identifying epitope on bovine prion protein bPrP(25–241) specifically recognized by a monoclonal antibody, 3E7 (mAb3E7) using XL-MS. After cross-linking with isotope-labeled crosslinkers disuccinimidyl suberate (DSS-d0/d12) and disuccinimidyl glutarate (DSG-d0/d6), the cross-linked immuno complex was digested by chymotrypsin and analyzed by liquid chromatography ESI-FTICR MS to achieve the high resolution and mass accuracy necessary to assign the cross-linked peptides without ambiguity. Other methods for epitope mapping, such as MS in conjunction with limited proteolysis and epitope excision, are also used [98, 99]. Lately HDX-MS has become the tool of choice for epitope/paratope mapping because of its fast turnaround time, less material requirement, peptide level resolution and relative ease in data analysis. HDX-MS is capable of determining both linear and conformational epitope. To determine the epitope/paratope, HDX rate of each backbone amide hydrogen in the Ag-Ab complex is compared to that in the free antigen or antibody. Those residues with slower backbone H/D exchange rate in the complex are due to additional protections caused by the binding, and therefore considered as the binding interface. Epitope mapping of a mAb raised against thrombin using offexchange of deuterated Ag-Ab complex and subsequent pepsin digestion and matrix assisted laser desorption/ionization MS was reported by Baerga-Ortiz et al. [100]. Using HDX-MS along with other orthogonal techniques, a group from Eli Lilly showed Hu007 (a humanized IgG1 mAb) binds to a region located in the open end of the β -barrel structure of IL-1β and blocks binding of IL-1β to its receptor [101]. The introduction of automation in HDX reaction and faster data analysis resulted in an increased application of HDX-MS in epitope mapping in the first half of this decade. However, reducing spectral complexity arising from large Ag-Ab complex and confidently identifying proteolytic peptides for desired sequence coverage was a challenge in earlier years when high-resolution mass spectrometers were not widely available. Coales et al. reported mapping the epitope of a small model protein (cytochrome c) by immobilizing the antibody to reduce the spectral complexity arising due to Ag-Ab complex [102], so that they were able to determine the epitope using a low-resolution instrument. The spectral complexity issue is mostly resolved when high-resolution MS instruments, such as Orbitrap, FTICR, or high-resolution TOF, are used. Alan Marshall’s group, using multiple proteolytic enzymes (pepsin and fungal protease XIII) and carefully optimizing automated HDX setup coupled to FTICR-MS, achieved more than 90% 14

sequence coverage and epitope identification of a large antigen (95kDa) - antibody complex. The identified epitope is consistent with mutagenesis, molecular modeling and electron microscopy studies [103]. With the advance of modern high-resolution MS, using HDX to identify epitope without immobilization of antibody has become routine and there have been many reports in the literature. Examples include epitope identification in factor H binding protein of Neisseria meningitides (Figure 3) [104, 105], chaperonin protein GroEL from Francisella tularensis [106], food allergen – almond Pru du 6 [107] and cashew Ana o 2 [108], pollen allergen Bet v 1[109], Notch3 negative regulatory region for activating and inhibiting antibodies [110, 111] and target epitope determination for anti-coagulation factor VIII antibodies [112, 113]. Up to date, epitope mapping using HDX-MS has been one of the most important applications in the therapeutic protein discovery. Comparing to the epitope determined by X-ray crystallography or scanning mutagenesis, an epitope determined by HDX-MS usually covers more residues. The reason is first due to the peptide level spatial resolution of the HDX-MS technique. More importantly, it is also due to the fact that HDX measures changes in H/D exchange rate instead of binding. Binding of an antigen to the antibody usually makes the binding site less flexible. As a result, in addition to the residues directly involved in the binding, H/D exchange rates of nearby residues often become slower due to this domain stabilization in the complex. Occasionally, binding of the antibody may cause some long-range allosteric effect that makes data interpretation more complicated. Since the residues involved in binding are on the surface of the antigen, they usually have very fast exchange rate in the free antigen. When slow-exchanging amide hydrogens in the free antigen become even slower in the complex, they are most likely not directly involved in binding but rather caused by domain stabilization or long range allosteric effect. Shorten the deuterium labeling time helps to reduce this effect [114] by distinguishing fast-exchanging amide hydrogens. Pandit et al. demonstrated that the combination of HDX-MS and computational docking (HDXDOCK) can improve the resolution of epitope mapping. They identified epitopes of cytochrome c-E8, IL13-CNTO607, and IL-17A-CAT-2200 interactions which are in good agreement with previous co-crystal structures. Next, the HDX-MS data was used as constraints during computational docking, resulting in more tightly clustered docking poses than stand-alone docking for all Ag-Ab interactions examined. This approach improved docking results significantly for the cytochrome c-E8 interaction [115]. Lim et al. used HDX-MS for both epitope and paratope mapping to study the interface of a dengue virus-neutralizing antibody, 2D22, with its target epitope in dengue virus strain DENV2 [116]. The epitope and paratope maps of 2D22-DENV2 complexes showed temperature-specific alterations in conformations and highlighted a heavy chain-mediated mode of binding of 2D22 at higher temperature. The paratope mapping revealed that the heavy chain interactions are primary determinants for stable 15

binding of 2D22 to DENV2, and consequently this interaction is maintained independent of the temperature-dependent expansion of DENV2. In recent years, FPOP has been used by Gross lab for epitope mapping. They reported the conformational epitope mapping of the serine protease thrombin [117] and solution binding interface of vascular endothelial growth factor against a fragment antigen binding region of an antibody [118]. FPOP offers several advantages over HDX-MS such as high speed irreversible labeling and side chain specificity. However, it involves complicated and time-consuming data analysis. Epitope information may also be obtained by differential chemical modification and MS [119]. The underlying principle is the differential surface-exposure and reagent-accessibility of a protein antigen in the presence and absence of an antibody that determines the rate and the extent of chemical modification of amino acid residues on the antigen.

3.1.3

Impact of protein conjugation

With the advancement of protein engineering and market expansion of biologics drug, the demand for hybrid novel modalities is also increasing. Antibody drug conjugates (ADCs) are combination products where a tumor-recognizing monoclonal antibody (mAb) is tagged with a small molecule drug to deliver a chemotherapeutic agent directly to cancer cells. To develop ADCs, it is critical that the choice of linker, conjugation site and the conjugation process do not interfere with the conformation and hence functional attributes of the mAb itself. To this end, Pan et al. [120] demonstrated, using a HDX-MS based method, that ADCs and mAb showed very similar conformational profile. A minor increase in HDX kinetics was due to the absence of intact interchain disulfide bonds and not because of the presence of pay load on the alkylated interchain cysteine residues. They followed this up with another HDX-MS study to demonstrate that site-specific drug conjugation at the engineered Cys residue at the position 239 of HC does not impact the structural integrity of antibodies [121]. They also demonstrated, in a separate study using sitespecific carboxyl group footprinting (a covalent labeling approach with GEE tags), that the conjugation did not alter the side-chain conformation [122]. Conjugation of polyethylene glycol (PEG) to a protein drug is often used to extend drug circulating life and stability. Using HDX-MS, Wei et al. studied the conformational changes of granulocyte colony stimulating factor (G-CSF) upon PEGylation [123]. The results indicated that although statistically significant differences in deuterium incorporation were induced by PEGylation of G-CSF, the overall changes were quite small. They concluded that PEGylation did not result in gross conformational rearrangement of G-CSF.

16

3.1.4

Understanding heterogeneity

IgG2, one of the major class of antibody with therapeutic applications, exhibits disulfide bond heterogeneity. Three distinct isoforms, IgG2-A, B and A/B have been identified and isolated using various biophysical and purification techniques [124-126]. Reversed-phase LC-UV-MS analysis revealed three main chromatographic peaks with the same mass of intact IgG2 antibody, but with dramatically different electrospray ionization charge distribution, when an IgG2 antibody was unfolded in the high temperature (75°C), high acidity (0.1% TFA, pH 2), and high organic (30% propyl alcohol) environment of the HPLC column [125]. The most compact B-form eluted first with fewer charges due to the most compact structure restricted by the disulfide bonds linking both Fab arms to the hinge.

A/B-form

possessing one flexible Fab arm and one attached to the hinge eluted next. A-form with most flexible Fab arms eluted latest and possessed largest numbers of charges, in agreement with the most open structure [125]. Besides understanding the conformational differences between these isoforms, it is critical to have a rapid detection and characterization method to facilitate the rapid progression of the molecule from discovery to commercial stage. To this end, Bagal et al. [127] developed a method where they leveraged the travelling wave ion mobility feature of the hybrid ion mobility quadrupole TOF MS (Synapt, Waters Inc., Milford, MA) and separated the B and A isoform of native IgG2 due to their different collision cross sections in gas phase. Their work demonstrated the ability of ion mobility as a shape selective separation methodology to detect disulfide heterogeneity in a mAb sample. Gross lab [128] used a combination of MS approaches, namely, ion mobility, top down MS sequencing and FPOP to characterize the disulfide heterogeneity in an IgG2 sample. They proposed that the combination MS approach is sufficiently advantageous and information rich and may become standard practice for characterizing novel biologics and biosimilars, minimizing the need for animal and human testing. Zhang et al. [129] reported the first detailed conformational analysis of three different disulfide isoforms using HDX-MS. The findings also suggest that the B isoform is more protected and more conformationally compact compared to the A isoform. The protection factor plots showed that the protected regions in the B isoform are located on the Fab domains, close to the hinge, centered on the side where the two Fab arms facing each other in spatial proximity. They proposed that in the more solventprotected B isoform, the two Fab arms are brought into contact by the non-classical disulfide bonds, resulting in a more compact global structure (Figure 4). Yan et al. [130] observed a mAb size variant consisting an extra light chain. To understand the mechanism of formation of this size variant, they used HDX-MS to characterize this variant and found that the residues near the N-terminus of the light chain (30-50) bind to the residues near the N-terminus of

17

the heavy chain (38-57). The information is helpful for construct design and process development of antibody-based biopharmaceuticals. Covalent labeling of surface-exposed residues can also be used to assess positive charge patch, when lysine and arginine side chains were targeted for labeling. Zhang et al. [131] has used sulfosuccinimidyl acetate and p-hydroxyphenylglyoxal as labeling reagents to label lysine and arginine side chains, respectively. LC-MS/MS peptide mapping data of a labeled mAb revealed the positive charge patch responsible for an anomalous charge variant observed in cation-exchange chromatography.

3.1.5 Molecular engineering

A successful biopharmaceutical drug candidate not only has a good efficacy and safety profile, it should also possess good physicochemical and pharmacokinetic properties. A molecular engineering process is often required to optimize a drug candidate for ideal physicochemical and pharmacokinetic properties. Geoghegana et al. applied the understanding from a HDX study to guide an IgG1 mAb variable domain re-engineering and thereby reduce the reversible self-association properties and viscosity [132]. The HDX study provided the molecular level understanding of the regions that are involved in the selfinteraction. The rational mutation strategies were built on those findings to reduce the surface hydrophobicity. As a result, the engineered mutations significantly reduced the reversible self-association and viscosity of the antibody, while maintaining the overall stability. To the same end, Dobson et al. described the similar strategy to improve the biophysical properties and pharmacokinetic profile of an anti-nerve growth factor antibody, MEDI1912 [133]. The antibody candidate has picomolar binding affinity to its target, but exhibits strong self-association, as well as the resulting poor pharmacokinetic profile. They performed HDX and cross-linking mass spectrometry to map the interface responsible for aberrant self-association. Furthermore, they applied in-silico tools based on structural MS findings to pinpoint the residues responsible for the self-association. From there, they were able to design three amino acid substitutions to enhance its biophysical properties and serum persistence, without compromising affinity and potency.

3.2

Cell Line, Cell Culture and Purification Development

After a therapeutic protein drug candidate is selected and optimized, a cell line is developed to express the candidate, and then the upstream cell culture process and downstream purification process must be created and optimized to produce the drug substance with a desired quality profile. To know what quality profile is desirable, MS-based conformational analysis has been used to understand the impact of different 18

quality attributes to the HOS of the protein. Additionally, MS-based method has also been used for HOS comparability/biosimilarity studies.

3.2.1

Impact of quality attributes

Recombinant biopharmaceuticals have an inherent degree of heterogeneity due to the complex biosynthetic processes used by living organisms to produce them. These quality attributes include but not limited to sequence variants, post-translational modifications such as glycosylation, hydroxylation, disulfide isoforms, etc. Heterogeneity also arises from non-enzymatic modifications such as oxidation, deamidation, isomerization, hydrolysis, aggregation, etc. during manufacturing, distribution and longterm storage. Some of the modifications may have no impact on safety and efficacy of the drug. Others, however, have severe impact on structure, antigen binding, Fc effector function, safety and hence the quality of the drug. During drug development, it is very important to understand which of these attributes are critical in terms of safety and efficacy. Impacts of several quality attributes to the conformation of the protein therapeutics have been studied with MS-based methods as described below.

Glycosylation Antibody glycosylation plays a critical role in the stability [134, 135], pharmacokinetics [136, 137] and antibody dependent cellular cytotoxicity [138-140]. It is a key contributor to molecular heterogeneity of IgG molecules and requires close monitoring throughout drug development process. The abundance of each glycoform varies greatly between cell lines, clones and production conditions [141-145]. The most well studied glycans in mAb molecules are the N-linked glycans at the CH2 domain (Asn297). Houde et al. [146] first carried out a comparative conformational analysis of glycosylated and deglycosylated IgG1 using HDX-MS. They found two regions in CH2, 236-242 and 242-253, became more flexible in deglycosylated form. These regions are also responsible for FcγRIII binding to antibody Fc [147]. In a follow-up HDX-MS study they observed no conformational changes when fucose was removed from the glycan. Extensive galactosylation, however, caused increased protection to some nearby residues, likely due to interactions between galactose and Lys-247 [148]. Jensen et al. [149] showed that deglycosylation also influenced the FcRn (neonatal Fc receptor) binding to IgG1 Fc region. They observed less conformational stabilization in 243-260 region in CH2 of deglycosylated IgG1 compare to glycosylated IgG1 in the presence of FcRn. Heck and coworkers [150] demonstrated mutations of Y407 in the CH3 domain of IgG1 and IgG4 significantly increased the sialylation, galactosylation, and branching of the N-linked glycans in the CH2 domain. Comparative HDX-MS of wild type and mutated Y407E IgG1 revealed a more exposed CH3–CH3 dimerization interface. They also 19

identified allosteric structural effects in the CH2 domain and in the CH2–CH3 interface. They proposed that these findings not only alter our understanding of antibody structure, but also reveals possibilities for obtaining recombinant IgG with glycosylation tailored for clinical applications. Using HDX-MS and limited proteolysis under a native-like condition, Fang et al. [151] carried out a systematic study to determine the impact of different glycoforms on the molecular conformation and stability of recombinant IgG1 and IgG2 molecules expressed from Chinese hamster ovary (CHO) cells. The protection factor plot created from MassAnalyzer clearly showed that IgG2 molecules containing high-mannose glycans are more flexible in the CH2 domain near residue Phe-243 and residues 273-306. Deuterium uptake curves of the glycan-containing peptides also demonstrated that both high-mannose and hybrid glycans increased the conformational flexibility in the CH2 domain and contributed to the decrease of the overall thermal stability, in agreement with previous structural studies of glycan influence on IgG structure. IgG molecules containing sialylated glycans in the CH2 domain exhibited similar proteolytic degradation behavior as high-mannose glycans, suggesting decreased CH2-domain stability compared to shorter complex glycans, likely resulting from steric effect that decreased the glycan-CH2 domain interaction (Figure 5).

Chemical modifications In addition to post-translational modifications occurring intracellularly, some non-enzymatic chemical modifications also occur during protein production and storage. Methionine oxidation is one of the most frequently observed modifications during mAb production and storage. Oxidation of two conserved methionine residues, Met-252 and Met-428, have been reported to decrease mAb thermal stability [152], protein A binding [153, 154], FcRn binding [154] and circulation half-life [155]. All these studies have established the correlations between the observed chemical modifications and corresponding effects on mAb stability, aggregation rate and biological functions. Several studies using MS-based methods have shed light on the conformational changes associated with these modifications and corresponding functional consequences from a HOS perspective. Using Native MS in combination with other biophysical techniques, Haberger et al. [156] developed a method to quantify and correlate Fc CH2 Met oxidation level with IgG1-huFcRn functional interaction. With careful optimization of cone voltage and collision energy in a Q-TOF Ultima MS system, they were able to resolve the FcRn, mAb and the complex. Their findings suggest that CH2 methionine oxidation results in a stepwise decrease of mAb3/huFcRn receptor complex formation. Quantitative correlation is only observed for doubly oxidized molecule, not for mono oxidized IgG. This study shows how MSbased HOS analysis represents a powerful tool to determine critical quality attributes (CQA). Other MS

20

based peptide mapping study also confirmed the importance of double oxidation of CH2 methionine but not the CH3 methionine residue in FcRn binding [153, 157, 158]. In 2009, Houde et al. [148] first reported the destabilization effect on mAb CH2 region 243-247 (within peptide fragment FLFPPKPKDTLMI) upon oxidation of Met252 and Met428 in an HDX-MS study. They proposed that the electrostatic repulsion due to double oxidation on these two residues causes a structural destabilization in the peptide backbone near 243-247 region. However, no effect was observed near Met-429, most likely due to its rigid antiparallel β-sheet structure. A similar observation was made by Bukitt et al. [159] when they compared several mAbs before and after accelerated oxidative stress. Taking four different mAbs with diverse sequence, Zhang et al. [160] carried out a detailed HDX-MS and homology modeling study to decipher the conformational changes upon Fc domain methionine oxidation. They found a similar trend as Houde et al. [148] and also observed more severe destabilization in CH2 due to oxidation in aglycosylated mAb compare to glycosylated sample. Thermal stability (as indicated by a lower thermal melting temperature, Tm, for CH2 domain) and aggregation propensity (as indicated by an increased formation of high molecular weight species) were also found to correlate well with the HDX-MS data. In the same report, they did not observe any significant conformational change upon Asn deamidation or Asp isomerization. However, they observed structural rigidity around Asp-55 and Asp-104 upon succinimide formation from Asp at mild acidic condition. Based on a previous finding that a marker peptide FLFPPKPKDTL in the CH2 region undergoes conformational changes upon Met oxidation, a group from Roche [161] developed a HDX-MS method to monitor structural changes due to chemical modifications at sensitivity levels realistic to the requirements of biopharmaceutical research and development (typically between 1 and 5% at peptide level). They first carried out a comparative HDX-MS time course experiment between 0% oxidized and 100% oxidized mAb samples, and identified the two marker peptides encompassing the FLFPPKPKDTL region, which underwent maximum deuterium uptake difference at 1-min time point. Next, in a targeted HDX-MS experiment at 1-min time point only, they compared nonoxidized mAb reference material (0%) with the mAb samples containing 1%, 5%, 10%, 20%, 40%, 60%, 80%, and 100% oxidized mAb spiked in. They were able to detect statistically significant difference (determined by Student’s t-test, p < 0.05 or p < 0.02) in deuterium uptake of the two marker peptides at the lowest spiked oxidized mAb content of 1%. This method demonstrated the sensitivity of HDX-MS workflow and its ability to monitor minor changes in mAb composition within a relatively short sample preparation, analysis and data processing time for potential CQA assessment.

Charge variants

21

Protein therapeutics are traditionally analyzed by charge-based chromatographic techniques such as ionexchange chromatography (IEX), in which charge variants are separated. Tang et al. [162] reported the use of HDX-MS to study the conformation of four different IgG1 charge variants. The IgG1, produced from an NS0 cell line, exhibited charge heterogeneity due to the presence of variants formed by Cterminal lysine truncation, deamidation, N-terminal Gln cyclization, and sialylation in the carbohydrate moiety. They isolated four different charge variants by cation exchange chromatography and characterized them using HDX-MS time course and SUPREX (stability of unpurified proteins from rates of HDX) techniques. The SUPREX technique assesses the unfolding profile and relative stability of the charge variants by measuring the exchange properties of globally protected amide protons in the presence of a chemical denaturant. These combined HDX studies, evaluating both solution dynamics and unfolding profile, demonstrated that all four charge variants were not significantly different in conformation, solution dynamics, chemical denaturant-induced unfolding profile and stability. It also proved that modifications listed above have little impact on the structure of the IgG1 molecule.

3.2.2

Comparability/biosimilarity study

Proteins are large complex molecules having distinct three-dimensional HOSs. These HOSs play a key role in the biological function of a protein. Due to their complex nature of production, biopharmaceuticals are susceptible to various modifications that influence their HOS and hence the desired function. During the development and manufacturing life cycle of a protein therapeutic product, a change in the manufacturing process is often necessary, such as when a scale-up is required for a large-scale clinical study. With a process change, a comparability study must be performed to establish consistency of the drug substance before and after the change, not only in their primary structure, but also in HOS. Even with the same manufacturing process, batch-to-batch consistency in HOS must be demonstrated. Additionally, development of biosimilar drugs requires a biosimilarity study to establish structural consistency between the biosimilar molecules and the innovator molecules. To establish consistency in HOS, biophysical techniques such as circular dichroism (CD), fluorescence, differential scanning calorimetry (DSC), isothermal titration calorimetry (ITC), analytical ultracentrifugation (AUC) and Fourier transform infrared spectroscopy (FTIR), etc. are generally used. None of these techniques, however, offers spatially resolved structural information. Multi-dimensional nuclear magnetic resonance spectroscopy (NMR), although capable of providing detailed structural information, is limited to only small proteins [163-165]. In recent years, HDX-MS has emerged as a sensitive and robust tool for comparability studies. We will discuss the application of HDX-MS for comparability studies of different production batches, biosimilars, and charge states. 22

Gross lab [166] first reported the use of HDX-MS to compare and differentiate the conformational states of four different insulin types (naturally produced bovine, procine and recombinantly produced human and its LysPro analog) used for treating insulin dependent diabetes. In this report they identified distinct HDX signatures on specific insulin molecules reflecting their unique conformation and folding and proposed this method for batch-to-batch variability testing. Nakazawa et al. [167] compared HDXMS profiles of intact recombinant insulin with several rapid acting analogs (lispro and glulisine), longacting analog (glargine) and two intermediate-acting preparations (protamine-containing formulations). They observed distinct HDX signature that correlates with pharmacokinetic behavior. In a follow up study [168] they studied human insulin and lispro with bottom-up HDX-MS and mapped, at peptide level, two regions (A3–6 and B22–24) in lispro analog exhibiting differences in deuterium uptake compared to the wild type. The differences observed in A3–6 and B22–24 regions are consistent with their known involvement in hexamer and dimer formation, respectively. Using interferon β as a model protein, Houde and coworker showed the application and usefulness of HDX-MS as a method for comparability studies of biotherapeutics between different batches with varying degree of modifications and storage conditions [42, 169, 170]. They also reported how HDX-MS can be used to demonstrate the conformational comparability between factor IX–Fc fusion protein (rFIX-Fc), factor IX (rFIX), and purified Fc fragment [171]. They showed that the HOS as detected by HDX-MS is highly comparable between individual rFIX and FIX region of rFIX-Fc, both in the absence and presence of calcium, a prerequisite for FIX function. They also established the conformational comparability between Fc and Fc region of rFIX–Fc. This study confirmed that fusing an IgG1 Fc to rFIX does not significantly alter the HOS of FIX or Fc, Ca binding to FIX, or Fc functionality. For biosimilar drugs, a thorough physiochemical and functional comparability exercise is key to establish consistency between the biosimilar and innovator drug and therefore gain regulatory approval. To this end, a group from Sandoz pharmaceutical used a battery of biophysical techniques, including HDX-MS, peptide mapping and functional assays to characterize a proposed biosimilar (GP2013) with its originator rituximab [172]. Their data demonstrated GP2013 to be highly similar to originator rituximab at the level of primary and HOS, post-translational modifications and size variants, and indistinguishable from a functional perspective. Fang et al. also compared the HOS of Remicade and biosimilar Inflectra by HDX-MS, and found no structural difference between the two products, except for very minor difference in the CH2 domain, likely caused by the slight difference in the glycan profile [173]. Pan et al. [174] developed a top down HDX-MS method by combining HDX with subzero temperature chromatography and electron transfer dissociation on the Orbitrap mass spectrometer. They were able to detect HDX information for 6 IgG domains on Herceptin, which included the antigen 23

binding sites. Ligand induced structural changes were found only in the variable regions. Additionally, global glycosylation profile of antibodies and HDX property of the glycoforms were also determined by accurate intact mass measurements. Because of minimal sample manipulation, fast workflow, very low level of back exchange, and simple data analysis, they proposed the suitability of this workflow for fast comparative structural evaluation of intact antibodies.

3.3 Formulation development During development of a potential protein therapeutics candidate, it is critical that the candidate molecule is formulated in a stable drug form suitable for storage, transport, and eventually dosing in clinical and commercial phase. In recent years, liquid formulation of therapeutic proteins and antibodies at 4°C for 2 years and several days at room temperature is highly preferred as compared to lyophilized and frozen formulations. During storage and transport, unfolding, aggregation, degradation or chemical modification of the molecule not only effect potency and efficacy but may also create fatal toxic and immunological side effect to patients [175-177]. Reducing degradation and instability, most notably aggregation, is a prime concern at formulation development phase. Moreover, biopharmaceuticals undergo many process challenges during their life cycle. Therefore, to establish commercial suitability, stability studies are typically performed at accelerated stressed conditions (temperature, pH, light), multiple freeze thaw (F/T) cycle, pH jump and drop shock, etc. To achieve a stable biopharmaceutical, a formulation development effort is undertaken to find an optimized matrix buffer and excipient composition (formulation buffer system) to reduce aggregation, increase stability, and meet the target product quality profile. MS-based technologies play a critical role in this juncture and we will discuss how MS-based technologies are implemented in characterizing protein HOS, understanding conformational stability and unfolding/folding behavior, etc. to help development of a stable formulation.

3.3.1

Conformational stability

When a protein domain is tightly folded, it is less likely to degrade or be modified. Therefore, understanding the conformational stability of different regions of the protein molecule is important in determining the root-cause of protein degradation, thereby facilitating design of better excipient for more stable conformation.

24

Determining conformational stability can be achieved by exposing the protein in increasing level of denaturant, and then following the unfolding process of the protein. HDX-MS, due to its spatial resolution, can be conveniently applied for this purpose. In a work performed by Zhang et al. [50], an IgG1 was exposed to different concentrations of guanidine during an HDX study. The resulting protection profile indicated that with increasing guanidine concentration, the CH2 domain unfolds first, followed by the variable domain. The CH3 domain, being the most stable domain in the IgG structure, does not start unfolding till 4 M guanidine. Because high concentration of guanidine may affect the intrinsic chemical exchange rate, a tetrapeptide, PPPI, was used to correct the effect of intrinsic HDX rate, due to its relatively slow exchange rate. Similar conclusion was also obtained using a partial reduction approach at different concentration of denaturant [178]. HDX-MS can also be used to study the conformational stability of lyophilized proteins in solid state [40, 41]. By exposing the lyophilized protein product to D2O vapor at a controlled vapor pressure in a desiccator, Topp and coworkers has labeled solid-state protein with gas-phase D2O and used bottom-up HDX-MS to study the conformational stability of recombinant proteins lyophilized with different excipients [43, 179]. It was found that the HDX data correlate strongly with long-term stability, suggesting solid-state HDX-MS can be used as a predictor of long-term stability and degradation rate of lyophilized protein [180, 181], which often takes months to years to obtain by conventional techniques. In addition, because of its ability to routinely furnish spatially resolved dynamic information in solid-state protein, HDX-MS provides insight into the mechanism of conformational instability during the drying process [182] or long-term storage, helping design better excipient for lyophilization.

3.3.2

Dimer, oligomer and aggregate formation

Size exclusion chromatography (SEC) is routinely used to quantify amount of high-molecular weight species, including dimers, oligomers and aggregates in a protein therapeutics. Native electrospray-MS, when coupled to SEC using volatile aqueous mobile phase, provides additional edge to characterization of these high-molecular weight species. Although native MS does not provide detailed molecular structure information, its sensitivity, speed, selectivity and ability to simultaneously measure several species present in a mixture are clearly advantageous in comparison with traditional structural biology methods. Kükrer et al. [5] reported native ESI-TOF analysis of intact human mAb aggregates fractionated by SEC. They generated IgG aggregates (dimers, trimers, tetramers and high-molecular-weight oligomers) by pH stress, fractionated by SEC, dialyzed against ammonium acetate pH 6.0 and analyzed by native ESI-TOF MS. They successfully identified monomeric IgG as well as dimers, trimers and tetramers.

25

Recently a group from Roche [7] reported the development of ultra-pressure SEC separation combined with native ESI-MS for the simultaneous formation, identification and quantification of size variants in recombinant antibodies. They carried out the SEC separation in ammonium acetate buffer on a Dionex UltiMate 3000 RSLC system interfaced with NanoMate direct infusion system. NanoMate was installed on the modified Waters High Mass Q-TOF Ultima mass spectrometer system, enabling measurement of protein/protein complexes at higher m/z. This setup allowed them to study the variants formed during formulation and bioprocess development, and can thus be potentially transferred to quality control units for routine in-process control and release analytics. A temperature-controlled ESI source in front of a Q-TOF mass spectrometer allowed real-time monitoring of unfolding and association of proteins as a function of source temperature [183]. The charge distribution of two therapeutic proteins (human antithrombin and an IgG1 mAb) under heat-stress conditions was monitored in real time, providing evidence that relatively small-scale conformational changes in each system lead to protein oligomerization, followed by aggregation. The ability of the temperature-controlled

ESI

MS

to

monitor

both

the

conformational

changes

and

oligomerization/degradation complements the classic calorimetric methods, positioning itself as a powerful analytical tool for the thermostability studies of protein therapeutics. Although Native MS provides straight-forward detection and quantification of mAb oligomers and aggregates, it does not provide any spatial resolution as provided by HDX-MS for conformational analysis. Zhang et al. [37] studied the effect of freezing stress on Bevacizumab antibody structure in the presence of a cryoprotectant (Trehalose) and different denaturing conditions (guanidine hydrochloride) using a previously developed frozen state HDX-MS protocol [36]. Briefly, they diluted the protein solution into D2O buffer (90% final deuterium) followed by flash freezing in liquid N2 for 5 min. The frozen samples were incubated for pre-determined labeling time (at -10°C), thawed quickly (2 min) by adding twice the sample volume of ice cold low pH denaturing quench buffer and subjected to LC/MS analysis. They reported two different mechanisms of aggregate formation upon applied stress: Bevacizumab formed native aggregate under freeze-thaw stress while non-native aggregate under thermal stress. Using HDX-MS, disulfide mapping and other orthogonal biophysical methods (small angel X ray scattering, DSC, non-reducing SDS PAGE and SEC), Iacob et al. [184] carried out comparative HOS analysis of two different mAb monomers and their corresponding dimers occurred during normal storage and production process. They found that one of the mAbs forms dimer by noncovalent interactions (determined by non-reducing SDS PAGE) and didn’t show any HDX footprint when compared to the monomer. Based on these findings and the observed changes in CH2 region by DSC, they speculated that the amino acid side chains of the CH2 domain are involved in noncovalent dimer formation. The other 26

mAb, on the contrary, forms a heterogeneous mixture of covalently attached dimers as determined by non-reducing SDS PAGE, a single broad DSC transition and small angel X ray scattering data analysis. Based on disulfide mapping analysis and HDX perturbation data, they proposed that the dimerization proceeded through domain swapping, disulfide scrambling and surface interactions predominantly in hinge and Fc CH2 regions. In agreement with other reports [185, 186], they identified a particularly sensitive region in CH2 domain, which becomes more dynamic upon dimer formation and remains tightly packed otherwise due to the stabilizing effect of nearby glycan moiety. To demonstrate the importance of this particular CH2 region in mAb aggregation, Majumdar et al. [187] carried out comparative HDX-MS study of an IgG1 and a triple mutant YTE (M255Y/S257T/T259E), engineered for extended half-life in vivo. Although they observed subtle or no differences in local flexibility at the mutated site, several other segments (VH, CH1 and VL) including 244-254 in CH2, showed significantly increased flexibility. DSC analysis showed decreases in both thermal onset and unfolding temperatures for the CH2 domain of the YTE mutant. In addition, YTE mutant aggregated faster than wild type under accelerated stability conditions as measured by SEC analysis. Correlating these observations, the relatively lower physical stability of the YTE mutant and concomitant increased local flexibility of the 244–254 segment, they proposed this particular segment of CH2 domain as potential aggregation hotspot in IgG1 mAb. A recent HDX-MS study of an Fc fusion protein also indicated towards the same region undergoing conformational changes upon dimer formation [188]. A group from Genentech used the emerging hydroxyl radical oxidation via synchrotron irradiation technology to map the interface in mAb dimers [189]. They enriched the dimer using SEC, oxidized both the monomer and dimer by hydroxyl radicals generated by exposure of the aqueous solution to synchrotron X-rays in millisecond timescales and carried out comparative bottom up LC/MS to identify the regions protected from oxidation in dimer. The dimer interface was identified in the Fab region and they proposed two possible head-to-head dimer models. The same conclusion was also obtained when they used papain and FabRICATOR® enzyme to separate the Fab and Fc, followed by SEC analysis accompanied with MS characterization to identify the Fab/Fab orientation of the IgG1 dimer.

3.3.3

Viscosity

Biologics are mostly administered via intravenous (IV) or subcutaneous (SC) route. Due to high cost, longer administration time and limited patient compliance associated with IV, SC administration as a prefilled syringe and autoinjector is gaining momentum [190]. The major challenge associated with high concentration, low volume prefilled dosage form is the high viscosity and associated protein instability and tissue backpressure [191]. MS-based methods have been used to decipher the mechanism of high 27

viscosity in order to identify potential hot spots driving the self-association responsible for the highviscosity. The challenge of studying protein viscosity by HDX is the high concentration of protein required during deuterium labeling, which excludes the routine method of starting deuterium labeling by direct dilution. To solve the problem, Arora et al. [38] modified the typical bottom-up HDX workflow by carrying out lyophilization of mAb solutions and subsequent reconstitution with a deuterated labeling buffer. Using this method, they characterized the non-covalent intermolecular interactions at high protein concentration, termed as reversible self-association (RSA). MAbs at low (5 mg/mL) and high (60 mg/ml) concentration at an RSA-promoting solution were lyophilized and reconstituted with a deuterated labeling buffer. The HDX-MS data suggested CDR2H and CDR2L as the major interaction interface at high concentration sample with other regions (VH, CH2 and hinge with increased flexibility) showing distant dynamic coupling effect. Using similar methodology in a follow up study, the same group reported mAb RSA driven by Fab-Fc protein-protein interaction [192]. Although the above HDX method analyzed protein at relatively high concentration (60 mg/ml) using lyophilization and subsequent reconstitution in deuterated labeling buffer, practical limitations such as reconstitution time, introduction of undesirable protein stresses and varying viscosity might restrict its applicability. Houda et al. [39] recently proposed an improved dilution free dialysis based HDX-MS method to characterize mAb at more than 200 mg/mL concentration.

3.3.4 Protein-excipient interactions Formulation development is carried out to increase the stability of drug product during manufacturing, storage, transportation and administration. Excipients such as sugars, salts, detergents, surfactants, and amino acids are added to the solution to stabilize the protein. Protein-excipient interaction is an active field of research [193, 194]. Traditionally, the effect of excipients on long term drug stability is empirically determined. High-throughput screening technology has been used to find suitable excipients to stabilize the drug molecule [195, 196]. HDX-MS has emerged as a valuable tool for understanding protein-excipient interactions. It has advantages over traditional techniques, such as X-ray, NMR, FTIR for faster turn-around time and higher spatial resolution (compared to FTIR). Volkins/Weis lab played a pioneering role in HDX-MS characterization of protein-excipient interactions. They characterized the effect of commonly used Hofmeister series salt (sodium salt of sulfate, chloride and thiocyante) and excipients, such as sucrose and arginine, on IgG1 mAb local dynamics and correlated it with the long-term stability during storage [185, 186]. Although the effect of these agents on melting temperature and aggregation propensity correlated well, local dynamics detected 28

by HDX-MS was somewhat complicated. Sucrose and chloride salt showed an overall decreased dynamics across several regions. These are believed to be due to preferential exclusion mechanism where sugar is preferentially excluded from the protein surface. Additionally, to accommodate such thermodynamically unfavorable condition, protein molecules undergo compactness to reduce its surface area. Sodium sulfate, although having positive effect on the physical stability (increased Tm), increased the local flexibility in CH1, CL, VH and VL domain of mAb as detected by HDX-MS. Thus, its stabilizing mechanism on mAb, although not clear, is different form sucrose and chloride salt. Arginine and thiocyante salt, on the contrary, have destabilizing effect on physical stability (decreased Tm and Tonset values, higher aggregation formation propensity) and also increased the local flexibility of mAb across different regions. One important aspect of arginine and thiocyante salt is that both substantially increase the flexibility of a particular consensus segment of CH2 domain (FLFPPKPKDTL). They correlated the local flexibility of this particular region in CH2 domain as a predictor of overall physical stability of mAb. Besides mAb, excipient interactions with other therapeutic proteins are also characterized using HDX-MS. Zhang et al. [197] systematically characterized the conformational changes on GCSF upon sucrose and benzyl alcohol addition. They observed, under a physiological condition, sucrose globally protects the whole molecule from deuterium exchange while benzyl alcohol induces increased deuterium uptake of the regions within the α-helical bundle. One drawback of protein-excipient interaction studies is the effect of these agents on the intrinsic HDX rates, which may produce false interpretation. To overcome this challenge, Zhang et al. [197] used a previously reported [50] model peptide (PPPI) to monitor the intrinsic exchange rate across all conditions. To this end, David Weis group reported [198] using an unstructured reporter peptide YPI to monitor the differences in chemical exchange rates in different solutions. An empirical chemical exchange correction factor, determined by use of the HDX data from the reporter peptide in different solutions, was then applied to the HDX measured in peptides derived from the protein of interest in the same solutions. They validated the correction through simulation and in a proof-of-concept HDX-MS excipient screening experiment using an IgG4 monoclonal antibody. The other challenge of routine implementation of HDX in formulation development studies is the unfavorable effect of different excipients on liquid chromatography (e.g., poor separation) and mass spectral quality (e.g., suppressing ionization of desired species). Removal of these excipients before LCMS while keeping the back exchange low is a challenging task. Kasper Rand and co-worker [199] developed a solid phase extraction HDX-MS procedure (SPE-HDX-MS) capable of removing interfering pharmaceutical excipients and carrying out global HDX analysis (intact protein analysis without proteolysis) of biologics. Using a reverse phase trap column and careful adjusting wash buffer solution to 29

remove excipients, they achieved considerable improvement in mass spectral signal-to-noise ratio and acceptable back exchange rate. Due to its suitability for high throughput screening using automation, they proposed the application of this method for pre-formulation screening of drug candidates.

4

Conclusions and Future Outlook

MS-based conformational analysis has been successfully applied in different stages of biopharmaceutical development, as discussed extensively in section 3 and summarized in terms of references in Table 1. At early stages, native MS, HDX-MS, oxidative footprinting and chemical cross-linking are used to analyze antibody-antigen complexes, where the epitope/paratope information is fed into the drug design/optimization process and intellectual property creation via the uncovering of detailed structural information. These MS-based methods are also being used for understanding molecular heterogeneity, assessing impact of protein conjugation, and molecular engineering. Further downstream in the biopharmaceutical development process, HDX-MS in particular is playing a dominant role in comparability/biosimilarity studies, as well as evaluating the structural impact of different quality attributes, e.g., glycosylation, oxidation and charge variants. In 2014, 11 biologics license applications (BLAs) were approved by FDA. Among those, three BLAs, including one mAb, one protein, and one fusion protein, used HDX-MS for HOS analysis [200]. Although a relatively high percentage (27%), HDX is still considered as a “nice to have” addition to filing. HDX-MS is also playing an important role in determining the best formulation conditions for product stability at particular pH, buffer, excipient and preservative combination. MS-based methods provide valuable information regarding protein-excipient interactions, as well as protein-protein interactions that are important in understanding viscosity and oligomerization and aggregation processes. It is very clear from this review that HDX-MS is a more favored technique for conformational analysis in biopharmaceutical development. Bottom-up HDX-MS, since its development more than two decades ago [21], has become the method of choice for many industrial laboratories for epitope mapping, comparability study, etc. First, HDX-MS is rich in information content; with a labeling site at virtually every residue. Although it usually provides peptide level spatial resolution, the intermediate resolution is advantageous over other biophysical techniques in providing detailed HOS information. Secondly, HDXMS has advantage of automated sample preparation and data analysis, and fast turn-around time. Native MS, due to its simplicity in data interpretation, is also widely used to determine the stoichiometry of protein-protein interactions. Oxidative footprinting [72, 77] is gaining momentum due to its advantage of irreversible labeling. Its application, however, is limited due to difficulty in the labeling

30

procedure and data interpretation. Chemical cross-linking, although not extensively used in the industry, has great potential when information extraction from raw data is more automated and streamlined. Oxidative footprinting and cross-linking remain challenging and require special training and skills to implement in an industrial environment, as compared to, for example, HDX-MS, which is becoming more routine. This is mainly because robust automation in sample handling and data processing is not available. For a technique to be widely acceptable by non-expert users in industrial laboratories, increased automation and robustness is the key. HDX-MS comes a long way and become a method of choice in many industrial laboratories, strongly owing to the cumulative efforts of improving robustness through automation in data collection and analysis. Increased automation and robustness in other techniques, especially information-rich methods such as oxidative footprinting and chemical cross-linking, will significantly increase their influences in the industry. Another important point to consider is the throughput required to analyze a large number of samples. Most MS-based methods require proteolytic digestion to gain spatially resolved HOS information. This procedure is time-consuming, labor-intensive and error-prone. Further advancement of top-down mass spectrometry should simplify sample preparation routine, minimize artificial modifications and make itself more automated and user friendly. This will require, of cause, robust instrumentation and software for processing the complex top-down data. Comparing to conventional biophysical techniques such as calorimetry, circular dichroism, fluorescence, infrared and Raman spectroscopy, etc., MS-based methods generally offer more detailed structural information, and are more tolerable to impurities in the sample. Comparing to high-resolution techniques such as X-ray diffraction and NMR, the spatial resolution provided by MS-based methods is limited. This disadvantage of MS-based methods, however, is offset by their fast turn-around time, small sample requirement, and applicability to large protein complexes. Although antibodies remain on the top of the list of protein therapeutic modalities, new entities, namely, bispecific antibodies, bispecific T-cell engagers, and antibody-drug conjugates are entering the pipelines and require MS attention. In the future, with the maturation of these MS-based methodologies, we expect that they will be applied more frequently and broadly in drug development processes. These methods will help us understand the unfolding mechanism of different protein constructs so that the amino acid sequence can be designed and optimized for more stable protein products. They will help us understand local unfolding/folding process to facilitate design and discovery of better excipient. They will also help us understand the criticality of quality attributes to achieve better quality control over a biopharmaceutical's overall development.

31

Table 1. Examples of MS-based conformation analyses of protein therapeutics during different stages of biopharmaceutical drug development.

Ab-Ag complex Epitope mapping Drug discovery, design and optimization

Cell Line, Cell Culture and Purification Development

Formulation development

Impact of protein conjugation Understanding heterogeneity Molecular engineering Impact of quality attributes Comparability/ biosimilarity study Conformational stability Dimer, oligomer and aggregate formation Viscosity Protein-excipient interactions

Native and nonnative MS of intact protein [90-96]

Limited proteolysis

HDX-MS

Covalent labeling

Crosslinking

[98, 99]

[100-116]

[117119] [122]

[97]

[120, 121, 123] [129, 130]

[127, 128]

[128]

[132, 133] [156]

[151]

[183] [5, 7, 183]

[189]

[146, 148, 150, 151, 159-162] [42, 166-174] [40, 41, 43, 50, 179-182] [36, 37, 184, 187, 188] [38, 39, 192] [185, 186, 197, 199]

32

[133]

[189]

FIGURE LEGEND Figure 1. Protein therapeutics development process and applications of MS-based conformational analysis. Figure 2. Native MS to determine stoichiometry of Denosumab-RANKL complex. Electrospray ionization mass spectra of A, aglycosylated RANKL at 0.08 µg/µl; B, denosumab at 0.2 µg/µl; C, their mixture at molar ratio of 3:2 of denosumab:RANKL-trimer and concentrations of 0.1 µg/µl and 0.04 µg/µl in 50 mM ammonium acetate; D, magnified view of the assembly 3D2R, partial assembly (620-710 kDa) and larger assembly region from C; and E, deconvoluted ESI mass spectra of the multiply charged ions created by denosumab, RANKL and their assemblies. Adapted with permission from Arthur et al. [93] Copyright 2012 American Chemical Society. Figure 3. Epitope mapping of factor H binding protein (fHbp) by HDX-MS. Boxes show deuterium uptake curves over 30 min for fHbp fragments in the absence (red) or presence (blue) of mAb 12C1. Deuterium incorporation was reduced by mAb 12C1 for each fragment shown. The peptides protected by mAb 12C1 in the HDX-MS analysis are highlighted in red on the fHbp structure. Numbered spheres indicate the start/end of each affected fragment. Adapted with permission from Malito et al. [104] Figure 4. Difference in protection factors of the light chain (A) and heavy chain (B) between the IgG2-B and IgG2-A disulfide isoforms (covalent structures shown in C), as determined by HDX-MS. Regions with more protection in the IgG2-B isoform are shown in blue on the 3-D structure of IgG2-A (D), indicating that in the more solvent-protected B isoform, the two Fab arms are brought into contact by the non-classical disulfide bonds, resulting in a more compact global structure. Adapted with permission from Zhang et al. [129] Copyright 2015 American Chemical Society. Figure 5. Abundance of released glycopeptides (sequence EEQYNSTYR) with different glycoforms after treatment of an IgG1 mAb with trypsin for varying lengths of time under a native-like condition. The longest time points are from complete digestion of the mAb, representing an infinite digestion time. Complex glycans (A2… or A3…) are shown on the top and all other glycans are shown on the bottom. Complex asialylated glycans (A2G… or A3G…) are shown in green, hybrid glycans (A1…) are shown in black, high-mannose glycans (M…) are shown in red, sialylated glycans (A2S…) are shown in blue, and minimally glycosylated species, including unglycosylated as well glycans containing one (Gn) or two (GnF) residues, are shown in brown. When two structural isomers are present, the early eluting one is labeled (a) and the late eluting one is labeled (b). High-mannose glycans, as well as some hybrid and sialylated glycans were found to release early, suggesting that they do not stabilize the CH2 domain as other complex glycans. Adapted with permission from Fang et al. [151] Copyright 2016 American Chemical Society.

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REFERENCES

[1] L.L. Green, Antibody engineering via genetic engineering of the mouse: XenoMouse strains are a vehicle for the facile generation of therapeutic human monoclonal antibodies, J Immunol Methods 231(12) (1999) 11-23. [2] Z. Zhang, H. Pan, X. Chen, Mass spectrometry for structural characterization of therapeutic antibodies, Mass Spectrom Rev 28(1) (2009) 147-76. [3] D.L. Smith, Z. Zhang, Probing noncovalent structural features of proteins by mass spectrometry, Mass Spectrom Rev 13(5-6) (1994) 411-429. [4] A.J. Heck, Native mass spectrometry: a bridge between interactomics and structural biology, Nature methods 5(11) (2008) 927. [5] B. Kukrer, V. Filipe, E. van Duijn, P.T. Kasper, R.J. Vreeken, A.J. Heck, W. Jiskoot, Mass spectrometric analysis of intact human monoclonal antibody aggregates fractionated by size-exclusion chromatography, Pharm Res 27(10) (2010) 2197-204. [6] K. Muneeruddin, J.J. Thomas, P.A. Salinas, I.A. Kaltashov, Characterization of small protein aggregates and oligomers using size exclusion chromatography with online detection by native electrospray ionization mass spectrometry, Anal Chem 86(21) (2014) 10692-10699. [7] M. Haberger, M. Leiss, A.K. Heidenreich, O. Pester, G. Hafenmair, M. Hook, L. Bonnington, H. Wegele, M. Haindl, D. Reusch, P. Bulau, Rapid characterization of biotherapeutic proteins by size-exclusion chromatography coupled to native mass spectrometry, MAbs 8(2) (2016) 331-9. [8] K. Muneeruddin, M. Nazzaro, I.A. Kaltashov, Characterization of intact protein conjugates and biopharmaceuticals using ion-exchange chromatography with online detection by native electrospray ionization mass spectrometry and top-down tandem mass spectrometry, Anal Chem 87(19) (2015) 10138-45. [9] Y. Leblanc, C. Ramon, N. Bihoreau, G. Chevreux, Charge variants characterization of a monoclonal antibody by ion exchange chromatography coupled on-line to native mass spectrometry: Case study after a long-term storage at +5 degrees C, J Chromatogr B Analyt Technol Biomed Life Sci 1048 (2017) 130-139. [10] M. Guilhaus, D. Selby, V. Mlynski, Orthogonal acceleration time-of-flight mass spectrometry, Mass Spectrom Rev 19(2) (2000) 65-107. [11] J.C. May, J.A. McLean, Ion mobility-mass spectrometry: time-dispersive instrumentation, Anal Chem 87(3) (2015) 1422-1436. [12] Q. Hu, R.J. Noll, H. Li, A. Makarov, M. Hardman, R. Graham Cooks, The Orbitrap: a new mass spectrometer, J Mass Spectrom 40(4) (2005) 430-443. [13] R.J. Rose, E. Damoc, E. Denisov, A. Makarov, A.J. Heck, High-sensitivity Orbitrap mass analysis of intact macromolecular assemblies, Nat Methods 9(11) (2012) 1084-6. [14] J. Janis, S. Pasanen, J. Rouvinen, P. Vainiotalo, Characterization of the pH-dependent dissociation of a multimeric metalloprotein Streptomyces rubiginosus xylose isomerase by ESI FT-ICR mass spectrometry, J Mass Spectrom 43(10) (2008) 1376-80. [15] E.N. Kitova, P.I. Kitov, D.R. Bundle, J.S. Klassen, The observation of multivalent complexes of Shigalike toxin with globotriaoside and the determination of their stoichiometry by nanoelectrospray Fouriertransform ion cyclotron resonance mass spectrometry, Glycobiology 11(7) (2001) 605-11. [16] B.T. Ruotolo, J.L. Benesch, A.M. Sandercock, S.-J. Hyung, C.V. Robinson, Ion mobility–mass spectrometry analysis of large protein complexes, Nature protocols 3(7) (2008) 1139. [17] Y. Zhong, S.-J. Hyung, B.T. Ruotolo, Ion mobility–mass spectrometry for structural proteomics, Expert Rev Proteomics 9(1) (2012) 47-58. 34

[18] A. Hvidt, K. Linderstrom-Lang, Exchange of hydrogen atoms in insulin with deuterium atoms in aqueous solutions, Biochim Biophys Acta 14(4) (1954) 574-5. [19] A. Hvidt, K. Linderstrom-Lang, The kinetics of the deuterium exchange of insulin with D2O; an amendment, Biochim Biophys Acta 16(1) (1955) 168-9. [20] S.W. Englander, L. Mayne, Protein folding studied using hydrogen-exchange labeling and twodimensional NMR, Annu Rev Biophys Biomol Struct 21 (1992) 243-65. [21] Z. Zhang, D.L. Smith, Determination of amide hydrogen exchange by mass spectrometry: a new tool for protein structure elucidation, Protein Sci 2(4) (1993) 522-31. [22] Y. Deng, Z. Zhang, D.L. Smith, Comparison of continuous and pulsed labeling amide hydrogen exchange/mass spectrometry for studies of protein dynamics, J Am Soc Mass Spectrom 10(8) (1999) 67584. [23] B.M. Hossain, L. Konermann, Pulsed hydrogen/deuterium exchange MS/MS for studying the relationship between noncovalent protein complexes in solution and in the gas phase after electrospray ionization, Anal Chem 78(5) (2006) 1613-9. [24] K.D. Rand, T.J. Jørgensen, Development of a peptide probe for the occurrence of hydrogen (1H/2H) scrambling upon gas-phase fragmentation, Anal Chem 79(22) (2007) 8686-8693. [25] K.D. Rand, C.M. Adams, R.A. Zubarev, T.J. Jorgensen, Electron capture dissociation proceeds with a low degree of intramolecular migration of peptide amide hydrogens, J Am Chem Soc 130(4) (2008) 13419. [26] J. Pan, J. Han, C.H. Borchers, L. Konermann, Electron capture dissociation of electrosprayed protein ions for spatially resolved hydrogen exchange measurements, J Am Soc Mass Spectrom 130(35) (2008) 11574-11575. [27] K.D. Rand, M. Zehl, O.N. Jensen, T.J. Jørgensen, Protein hydrogen exchange measured at singleresidue resolution by electron transfer dissociation mass spectrometry, Anal Chem 81(14) (2009) 55775584. [28] J. Pan, J. Han, C.H. Borchers, L. Konermann, Hydrogen/deuterium exchange mass spectrometry with top-down electron capture dissociation for characterizing structural transitions of a 17 kDa protein, J Am Chem Soc 131(35) (2009) 12801-8. [29] G. Wang, R.R. Abzalimov, C.E. Bobst, I.A. Kaltashov, Conformer-specific characterization of nonnative protein states using hydrogen exchange and top-down mass spectrometry, Proc Natl Acad Sci U S A 110(50) (2013) 20087-92. [30] S.J. Eyles, T. Dresch, L.M. Gierasch, I.A. Kaltashov, Unfolding dynamics of a beta-sheet protein studied by mass spectrometry, J Mass Spectrom 34(12) (1999) 1289-95. [31] T. Rob, P. Liuni, P.K. Gill, S. Zhu, N. Balachandran, P.J. Berti, D.J. Wilson, Measuring dynamics in weakly structured regions of proteins using microfluidics-enabled subsecond H/D exchange mass spectrometry, Anal Chem 84(8) (2012) 3771-9. [32] D.A. Simmons, S.D. Dunn, L. Konermann, Conformational dynamics of partially denatured myoglobin studied by time-resolved electrospray mass spectrometry with online hydrogen-deuterium exchange, Biochemistry 42(19) (2003) 5896-905. [33] K. Dharmasiri, D.L. Smith, Mass spectrometric determination of isotopic exchange rates of amide hydrogens located on the surfaces of proteins, Anal Chem 68(14) (1996) 2340-4. [34] D. Goswami, S. Devarakonda, M.J. Chalmers, B.D. Pascal, B.M. Spiegelman, P.R. Griffin, Time window expansion for HDX analysis of an intrinsically disordered protein, J Am Soc Mass Spectrom 24(10) (2013) 1584-92. [35] S.J. Coales, S.Y. E, J.E. Lee, A. Ma, J.A. Morrow, Y. Hamuro, Expansion of time window for mass spectrometric measurement of amide hydrogen/deuterium exchange reactions, Rapid Commun Mass Spectrom 24(24) (2010) 3585-92.

35

[36] A. Zhang, W. Qi, S.K. Singh, E.J. Fernandez, A new approach to explore the impact of freeze-thaw cycling on protein structure: hydrogen/deuterium exchange mass spectrometry (HX-MS), Pharm Res 28(5) (2011) 1179-93. [37] A. Zhang, S.K. Singh, M.R. Shirts, S. Kumar, E.J. Fernandez, Distinct aggregation mechanisms of monoclonal antibody under thermal and freeze-thaw stresses revealed by hydrogen exchange, Pharm Res 29(1) (2012) 236-50. [38] J. Arora, J.M. Hickey, R. Majumdar, R. Esfandiary, S.M. Bishop, H.S. Samra, C.R. Middaugh, D.D. Weis, D.B. Volkin, Hydrogen exchange mass spectrometry reveals protein interfaces and distant dynamic coupling effects during the reversible self-association of an IgG1 monoclonal antibody, MAbs 7(3) (2015) 525-39. [39] D. Houde, Z.E. Nazari, G.M. Bou-Assaf, A.S. Weiskopf, K.D. Rand, Conformational Analysis of Proteins in Highly Concentrated Solutions by Dialysis-Coupled Hydrogen/Deuterium Exchange Mass Spectrometry, J Am Soc Mass Spectrom 27(4) (2016) 669-76. [40] Y. Li, T.D. Williams, R.L. Schowen, E.M. Topp, Characterizing protein structure in amorphous solids using hydrogen/deuterium exchange with mass spectrometry, Anal Biochem 366(1) (2007) 18-28. [41] Y. Li, T.D. Williams, R.L. Schowen, E.M. Topp, Trehalose and calcium exert site-specific effects on calmodulin conformation in amorphous solids, Biotechnol Bioeng 97(6) (2007) 1650-3. [42] C.E. Bobst, R.R. Abzalimov, D. Houde, M. Kloczewiak, R. Mhatre, S.A. Berkowitz, I.A. Kaltashov, Detection and characterization of altered conformations of protein pharmaceuticals using complementary mass spectrometry-based approaches, Anal Chem 80(19) (2008) 7473-81. [43] S. Sinha, Y. Li, T.D. Williams, E.M. Topp, Protein conformation in amorphous solids by FTIR and by hydrogen/deuterium exchange with mass spectrometry, Biophys J 95(12) (2008) 5951-61. [44] A.M. Sophocleous, J. Zhang, E.M. Topp, Localized hydration in lyophilized myoglobin by hydrogendeuterium exchange mass spectrometry. 1. Exchange mapping, Mol Pharm 9(4) (2012) 718-26. [45] V.L. Woods, Jr., Y. Hamuro, High resolution, high-throughput amide deuterium exchange-mass spectrometry (DXMS) determination of protein binding site structure and dynamics: utility in pharmaceutical design, J Cell Biochem Suppl Suppl 37 (2001) 89-98. [46] Y. Hamuro, S.J. Coales, M.R. Southern, J.F. Nemeth-Cawley, D.D. Stranz, P.R. Griffin, Rapid analysis of protein structure and dynamics by hydrogen/deuterium exchange mass spectrometry, J Biomol Tech 14(3) (2003) 171-82. [47] S. Kazazic, H.M. Zhang, T.M. Schaub, M.R. Emmett, C.L. Hendrickson, G.T. Blakney, A.G. Marshall, Automated data reduction for hydrogen/deuterium exchange experiments, enabled by high-resolution Fourier transform ion cyclotron resonance mass spectrometry, J Am Soc Mass Spectrom 21(4) (2010) 550-8. [48] M.J. Chalmers, S.A. Busby, B.D. Pascal, Y. He, C.L. Hendrickson, A.G. Marshall, P.R. Griffin, Probing protein ligand interactions by automated hydrogen/deuterium exchange mass spectrometry, Anal Chem 78(4) (2006) 1005-14. [49] D.J. Cummins, A. Espada, S.J. Novick, M. Molina-Martin, R.E. Stites, J.F. Espinosa, H. Broughton, D. Goswami, B.D. Pascal, J.A. Dodge, M.J. Chalmers, P.R. Griffin, Two-Site Evaluation of the Repeatability and Precision of an Automated Dual-Column Hydrogen/Deuterium Exchange Mass Spectrometry Platform, Anal Chem 88(12) (2016) 6607-14. [50] Z. Zhang, A. Zhang, G. Xiao, Improved protein hydrogen/deuterium exchange mass spectrometry platform with fully automated data processing, Anal Chem 84(11) (2012) 4942-9. [51] Z. Zhang, J. Fang, Extracting Information from Hydrogen Exchange Mass Spectrometry Data, in: D.D. Weis (Ed.), Hydrogen Exchange Mass Spectrometry of Proteins: Fundamentals, Methods, and Applications, John Wiley & Sons Ltd.2016, p. 107.

36

[52] M. Palmblad, J. Buijs, P. Hakansson, Automatic analysis of hydrogen/deuterium exchange mass spectra of peptides and proteins using calculations of isotopic distributions, J Am Soc Mass Spectrom 12(11) (2001) 1153-62. [53] Y. Hamuro, S.J. Coales, J.A. Morrow, K.S. Molnar, S.J. Tuske, M.R. Southern, P.R. Griffin, Hydrogen/deuterium-exchange (H/D-Ex) of PPARgamma LBD in the presence of various modulators, Protein Sci 15(8) (2006) 1883-92. [54] D.D. Weis, J.R. Engen, I.J. Kass, Semi-automated data processing of hydrogen exchange mass spectra using HX-Express, J Am Soc Mass Spectrom 17(12) (2006) 1700-3. [55] M. Hotchko, G.S. Anand, E.A. Komives, L.F. Ten Eyck, Automated extraction of backbone deuteration levels from amide H/2H mass spectrometry experiments, Protein Sci 15(3) (2006) 583-601. [56] G.W. Slysz, C.A. Baker, B.M. Bozsa, A. Dang, A.J. Percy, M. Bennett, D.C. Schriemer, Hydra: software for tailored processing of H/D exchange data from MS or tandem MS analyses, BMC Bioinformatics 10 (2009) 162. [57] X. Lou, M. Kirchner, B.Y. Renard, U. Kothe, S. Boppel, C. Graf, C.T. Lee, J.A. Steen, H. Steen, M.P. Mayer, F.A. Hamprecht, Deuteration distribution estimation with improved sequence coverage for HX/MS experiments, Bioinformatics 26(12) (2010) 1535-41. [58] Z.Y. Kan, L. Mayne, P.S. Chetty, S.W. Englander, ExMS: data analysis for HX-MS experiments, J Am Soc Mass Spectrom 22(11) (2011) 1906-15. [59] B.D. Pascal, M.J. Chalmers, S.A. Busby, P.R. Griffin, HD desktop: an integrated platform for the analysis and visualization of H/D exchange data, J Am Soc Mass Spectrom 20(4) (2009) 601-10. [60] B.D. Pascal, S. Willis, J.L. Lauer, R.R. Landgraf, G.M. West, D. Marciano, S. Novick, D. Goswami, M.J. Chalmers, P.R. Griffin, HDX workbench: software for the analysis of H/D exchange MS data, J Am Soc Mass Spectrom 23(9) (2012) 1512-21. [61] B.D. Pascal, M.J. Chalmers, S.A. Busby, C.C. Mader, M.R. Southern, N.F. Tsinoremas, P.R. Griffin, The Deuterator: software for the determination of backbone amide deuterium levels from H/D exchange MS data, BMC Bioinformatics 8 (2007) 156. [62] D.E. Miller, C.B. Prasannan, M.T. Villar, A.W. Fenton, A. Artigues, HDXFinder: automated analysis and data reporting of deuterium/hydrogen exchange mass spectrometry, J Am Soc Mass Spectrom 23(2) (2012) 425-9. [63] Z.Y. Kan, B.T. Walters, L. Mayne, S.W. Englander, Protein hydrogen exchange at residue resolution by proteolytic fragmentation mass spectrometry analysis, Proc Natl Acad Sci U S A 110(41) (2013) 16438-43. [64] K.M. Burns, M. Rey, C.A. Baker, D.C. Schriemer, Platform dependencies in bottom-up hydrogen/deuterium exchange mass spectrometry, Mol Cell Proteomics 12(2) (2013) 539-48. [65] P.G. Fajer, G.M. Bou-Assaf, A.G. Marshall, Improved sequence resolution by global analysis of overlapped peptides in hydrogen/deuterium exchange mass spectrometry, J Am Soc Mass Spectrom 23(7) (2012) 1202-8. [66] J. Zhang, P. Ramachandran, R. Kumar, M.L. Gross, H/D exchange centroid monitoring is insufficient to show differences in the behavior of protein states, J Am Soc Mass Spectrom 24(3) (2013) 450-3. [67] V.L. Mendoza, R.W. Vachet, Probing protein structure by amino acid-specific covalent labeling and mass spectrometry, Mass Spectrom Rev 28(5) (2009) 785-815. [68] H. Zhang, J. Wen, R.Y. Huang, R.E. Blankenship, M.L. Gross, Mass spectrometry-based carboxyl footprinting of proteins: method evaluation, Int J Mass Spectrom 312 (2012) 78-86. [69] L. Wang, M.R. Chance, Structural mass spectrometry of proteins using hydroxyl radical based protein footprinting, Anal Chem 83(19) (2011) 7234-7241. [70] G. Xu, M.R. Chance, Hydroxyl radical-mediated modification of proteins as probes for structural proteomics, Chem Rev 107(8) (2007) 3514-43. [71] S.D. Maleknia, M. Brenowitz, M.R. Chance, Millisecond radiolytic modification of peptides by synchrotron X-rays identified by mass spectrometry, Anal Chem 71(18) (1999) 3965-73. 37

[72] M.R. Chance, Unfolding of apomyoglobin examined by synchrotron footprinting, Biochem Biophys Res Commun 287(3) (2001) 614-21. [73] T.D. Tullius, B.A. Dombroski, Hydroxyl radical "footprinting": high-resolution information about DNAprotein contacts and application to lambda repressor and Cro protein, Proc Natl Acad Sci U S A 83(15) (1986) 5469-73. [74] B. Sclavi, S. Woodson, M. Sullivan, M. Chance, M. Brenowitz, Following the folding of RNA with timeresolved synchrotron X-ray footprinting, Methods Enzymol 295 (1998) 379-402. [75] J.J. Hayes, L. Kam, T.D. Tullius, Footprinting protein-DNA complexes with gamma-rays, Methods Enzymol 186 (1990) 545-9. [76] J.S. Sharp, J.M. Becker, R.L. Hettich, Analysis of protein solvent accessible surfaces by photochemical oxidation and mass spectrometry, Anal Chem 76(3) (2004) 672-83. [77] D.M. Hambly, M.L. Gross, Laser flash photolysis of hydrogen peroxide to oxidize protein solventaccessible residues on the microsecond timescale, J Am Soc Mass Spectrom 16(12) (2005) 2057-63. [78] T.T. Aye, T.Y. Low, S.K. Sze, Nanosecond laser-induced photochemical oxidation method for protein surface mapping with mass spectrometry, Anal Chem 77(18) (2005) 5814-22. [79] B.C. Gau, J.S. Sharp, D.L. Rempel, M.L. Gross, Fast photochemical oxidation of protein footprints faster than protein unfolding, Anal Chem 81(16) (2009) 6563-71. [80] J. Chen, W. Cui, D. Giblin, M.L. Gross, New protein footprinting: fast photochemical iodination combined with top-down and bottom-up mass spectrometry, J Am Soc Mass Spectrom 23(8) (2012) 1306-18. [81] M. Cheng, B. Zhang, W. Cui, M.L. Gross, Laser-Initiated Radical Trifluoromethylation of Peptides and Proteins: Application to Mass-Spectrometry-Based Protein Footprinting, Angew Chem Int Ed Engl 56(45) (2017) 14007-14010. [82] P. Kaur, J.G. Kiselar, M.R. Chance, Integrated algorithms for high-throughput examination of covalently labeled biomolecules by structural mass spectrometry, Anal Chem 81(19) (2009) 8141-9. [83] M.M. Young, N. Tang, J.C. Hempel, C.M. Oshiro, E.W. Taylor, I.D. Kuntz, B.W. Gibson, G. Dollinger, High throughput protein fold identification by using experimental constraints derived from intramolecular cross-links and mass spectrometry, Proc Natl Acad Sci U S A 97(11) (2000) 5802-6. [84] A. Leitner, M. Faini, F. Stengel, R. Aebersold, Crosslinking and Mass Spectrometry: An Integrated Technology to Understand the Structure and Function of Molecular Machines, Trends Biochem Sci 41(1) (2016) 20-32. [85] P.E. Kinnear, E.G. Tuddenham, Albinism with haemorrhagic diathesis: Hermansky-Pudlak syndrome, Br J Ophthalmol 69(12) (1985) 904-8. [86] P.d.L. A Fontana, B Spolaore, E Frare, P Picotti, M Zambonin, Probing protein structure by limited proteolysis, ACTA BIOCHIMICA POLONICA-ENGLISH 51 (2004) 299-322. [87] M. Monti, S. Principe, S. Giorgetti, P. Mangione, G. Merlini, A. Clark, V. Bellotti, A. Amoresano, P. Pucci, Topological investigation of amyloid fibrils obtained from beta2-microglobulin, Protein Sci 11(10) (2002) 2362-9. [88] F. Zappacosta, A. Pessi, E. Bianchi, S. Venturini, M. Sollazzo, A. Tramontano, G. Marino, P. Pucci, Probing the tertiary structure of proteins by limited proteolysis and mass spectrometry: the case of Minibody, Protein Sci 5(5) (1996) 802-813. [89] J. Villanueva, V. Villegas, E. Querol, F.X. Aviles, L. Serrano, Protein secondary structure and stability determined by combining exoproteolysis and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry, J Mass Spectrom 37(9) (2002) 974-84. [90] M.A. Tito, J. Miller, N. Walker, K.F. Griffin, E.D. Williamson, D. Despeyroux-Hill, R.W. Titball, C.V. Robinson, Probing molecular interactions in intact antibody: antigen complexes, an electrospray time-offlight mass spectrometry approach, Biophys J 81(6) (2001) 3503-9.

38

[91] M. Oda, S. Uchiyama, C.V. Robinson, K. Fukui, Y. Kobayashi, T. Azuma, Regional and segmental flexibility of antibodies in interaction with antigens of different size, FEBS J 273(7) (2006) 1476-87. [92] M. Oda, S. Uchiyama, M. Noda, Y. Nishi, M. Koga, K. Mayanagi, C.V. Robinson, K. Fukui, Y. Kobayashi, K. Morikawa, T. Azuma, Effects of antibody affinity and antigen valence on molecular forms of immune complexes, Mol Immunol 47(2-3) (2009) 357-64. [93] K.K. Arthur, J.P. Gabrielson, N. Hawkins, D. Anafi, J. Wypych, A. Nagi, J.K. Sullivan, P.V. Bondarenko, In vitro stoichiometry of complexes between the soluble RANK ligand and the monoclonal antibody denosumab, Biochemistry 51(3) (2012) 795-806. [94] C. Atmanene, E. Wagner-Rousset, M. Malissard, B. Chol, A. Robert, N. Corvaia, A. Van Dorsselaer, A. Beck, S. Sanglier-Cianferani, Extending mass spectrometry contribution to therapeutic monoclonal antibody lead optimization: characterization of immune complexes using noncovalent ESI-MS, Anal Chem 81(15) (2009) 6364-73. [95] F. Debaene, E. Wagner-Rousset, O. Colas, D. Ayoub, N. Corvaia, A. Van Dorsselaer, A. Beck, S. Cianferani, Time resolved native ion-mobility mass spectrometry to monitor dynamics of IgG4 Fab arm exchange and "bispecific" monoclonal antibody formation, Anal Chem 85(20) (2013) 9785-92. [96] A. Dyachenko, G. Wang, M. Belov, A. Makarov, R.N. de Jong, E.T. van den Bremer, P.W. Parren, A.J. Heck, Tandem Native Mass-Spectrometry on Antibody-Drug Conjugates and Submillion Da AntibodyAntigen Protein Assemblies on an Orbitrap EMR Equipped with a High-Mass Quadrupole Mass Selector, Anal Chem 87(12) (2015) 6095-102. [97] T. Pimenova, A. Nazabal, B. Roschitzki, J. Seebacher, O. Rinner, R. Zenobi, Epitope mapping on bovine prion protein using chemical cross-linking and mass spectrometry, J Mass Spectrom 43(2) (2008) 185-95. [98] C. Hager-Braun, H. Katinger, K.B. Tomer, The HIV-neutralizing monoclonal antibody 4E10 recognizes N-terminal sequences on the native antigen, J Immunol 176(12) (2006) 7471-81. [99] J.G. Kiselar, K.M. Downard, Direct identification of protein epitopes by mass spectrometry without immobilization of antibody and isolation of antibody-peptide complexes, Anal Chem 71(9) (1999) 1792801. [100] A. Baerga-Ortiz, C.A. Hughes, J.G. Mandell, E.A. Komives, Epitope mapping of a monoclonal antibody against human thrombin by H/D-exchange mass spectrometry reveals selection of a diverse sequence in a highly conserved protein, Protein Sci 11(6) (2002) 1300-8. [101] J. Lu, D.R. Witcher, M.A. White, X. Wang, L. Huang, R. Rathnachalam, J.M. Beals, S. Kuhstoss, IL1beta epitope mapping using site-directed mutagenesis and hydrogen-deuterium exchange mass spectrometry analysis, Biochemistry 44(33) (2005) 11106-14. [102] S.J. Coales, S.J. Tuske, J.C. Tomasso, Y. Hamuro, Epitope mapping by amide hydrogen/deuterium exchange coupled with immobilization of antibody, on-line proteolysis, liquid chromatography and mass spectrometry, Rapid Commun Mass Spectrom 23(5) (2009) 639-47. [103] Q. Zhang, L.N. Willison, P. Tripathi, S.K. Sathe, K.H. Roux, M.R. Emmett, G.T. Blakney, H.M. Zhang, A.G. Marshall, Epitope mapping of a 95 kDa antigen in complex with antibody by solution-phase amide backbone hydrogen/deuterium exchange monitored by Fourier transform ion cyclotron resonance mass spectrometry, Anal Chem 83(18) (2011) 7129-36. [104] E. Malito, A. Faleri, P. Lo Surdo, D. Veggi, G. Maruggi, E. Grassi, E. Cartocci, I. Bertoldi, A. Genovese, L. Santini, G. Romagnoli, E. Borgogni, S. Brier, C. Lo Passo, M. Domina, F. Castellino, F. Felici, S. van der Veen, S. Johnson, S.M. Lea, C.M. Tang, M. Pizza, S. Savino, N. Norais, R. Rappuoli, M.J. Bottomley, V. Masignani, Defining a protective epitope on factor H binding protein, a key meningococcal virulence factor and vaccine antigen, Proc Natl Acad Sci U S A 110(9) (2013) 3304-9. [105] A. Faleri, L. Santini, S. Brier, W. Pansegrau, P. Lo Surdo, M. Scarselli, F. Buricchi, G. Volpini, A. Genovese, S. van der Veen, S. Lea, C.M. Tang, S. Savino, M. Pizza, O. Finco, N. Norais, V. Masignani, Two cross-reactive monoclonal antibodies recognize overlapping epitopes on Neisseria meningitidis factor H binding protein but have different functional properties, FASEB J 28(4) (2014) 1644-53. 39

[106] Z. Lu, M.J. Rynkiewicz, G. Madico, S. Li, C.Y. Yang, H.M. Perkins, S.R. Sompuram, V. Kodela, T. Liu, T. Morris, D. Wang, M.I. Roche, B.A. Seaton, J. Sharon, B-cell epitopes in GroEL of Francisella tularensis, PLoS One 9(6) (2014) e99847. [107] L.N. Willison, Q. Zhang, M. Su, S.S. Teuber, S.K. Sathe, K.H. Roux, Conformational epitope mapping of Pru du 6, a major allergen from almond nut, Mol Immunol 55(3-4) (2013) 253-63. [108] Q. Zhang, K.A. Noble, Y. Mao, N.L. Young, S.K. Sathe, K.H. Roux, A.G. Marshall, Rapid screening for potential epitopes reactive with a polycolonal antibody by solution-phase H/D exchange monitored by FTICR mass spectrometry, J Am Soc Mass Spectrom 24(7) (2013) 1016-25. [109] S. Brier, M. Le Mignon, K. Jain, C. Lebrun, F. Peurois, C. Kellenberger, V. Bordas-Le Floch, L. Mascarell, E. Nony, P. Moingeon, Characterization of epitope specificities of reference antibodies used for the quantification of the birch pollen allergen Bet v 1, Allergy (2017). [110] K. Tiyanont, T.E. Wales, C.W. Siebel, J.R. Engen, S.C. Blacklow, Insights into Notch3 activation and inhibition mediated by antibodies directed against its negative regulatory region, J Mol Biol 425(17) (2013) 3192-204. [111] K. Tiyanont, T.E. Wales, M. Aste-Amezaga, J.C. Aster, J.R. Engen, S.C. Blacklow, Evidence for increased exposure of the Notch1 metalloprotease cleavage site upon conversion to an activated conformation, Structure 19(4) (2011) 546-54. [112] A.M. Sevy, J.F. Healey, W. Deng, P.C. Spiegel, S.L. Meeks, R. Li, Epitope mapping of inhibitory antibodies targeting the C2 domain of coagulation factor VIII by hydrogen-deuterium exchange mass spectrometry, J Thromb Haemost 11(12) (2013) 2128-36. [113] E. Bloem, M. van den Biggelaar, A. Wroblewska, J. Voorberg, J.H. Faber, M. Kjalke, H.R. Stennicke, K. Mertens, A.B. Meijer, Factor VIII C1 domain spikes 2092-2093 and 2158-2159 comprise regions that modulate cofactor function and cellular uptake, J Biol Chem 288(41) (2013) 29670-9. [114] B. Deng, S. Zhu, A.M. Macklin, J. Xu, C. Lento, A. Sljoka, D.J. Wilson, Suppressing allostery in epitope mapping experiments using millisecond hydrogen / deuterium exchange mass spectrometry, MAbs 9(8) (2017) 1327-1336. [115] D. Pandit, S.J. Tuske, S.J. Coales, S.Y. E, A. Liu, J.E. Lee, J.A. Morrow, J.F. Nemeth, Y. Hamuro, Mapping of discontinuous conformational epitopes by amide hydrogen/deuterium exchange mass spectrometry and computational docking, J Mol Recognit 25(3) (2012) 114-24. [116] X.X. Lim, A. Chandramohan, X.E. Lim, J.E. Crowe, Jr., S.M. Lok, G.S. Anand, Epitope and Paratope Mapping Reveals Temperature-Dependent Alterations in the Dengue-Antibody Interface, Structure 25(9) (2017) 1391-1402 e3. [117] L.M. Jones, B.S. J, A.C. J, M.L. Gross, Fast photochemical oxidation of proteins for epitope mapping, Anal Chem 83(20) (2011) 7657-61. [118] Y. Zhang, A.T. Wecksler, P. Molina, G. Deperalta, M.L. Gross, Mapping the Binding Interface of VEGF and a Monoclonal Antibody Fab-1 Fragment with Fast Photochemical Oxidation of Proteins (FPOP) and Mass Spectrometry, J Am Soc Mass Spectrom 28(5) (2017) 850-858. [119] S. Dhungana, M.B. Fessler, K.B. Tomer, Epitope mapping by differential chemical modification of antigens, Methods Mol Biol 524 (2009) 119-34. [120] L.Y. Pan, O. Salas-Solano, J.F. Valliere-Douglass, Conformation and dynamics of interchain cysteinelinked antibody-drug conjugates as revealed by hydrogen/deuterium exchange mass spectrometry, Anal Chem 86(5) (2014) 2657-64. [121] L.Y. Pan, O. Salas-Solano, J.F. Valliere-Douglass, Antibody structural integrity of site-specific antibody-drug conjugates investigated by hydrogen/deuterium exchange mass spectrometry, Anal Chem 87(11) (2015) 5669-76. [122] L.Y. Pan, O. Salas-Solano, J.F. Valliere-Douglass, Localized conformational interrogation of antibody and antibody-drug conjugates by site-specific carboxyl group footprinting, MAbs 9(2) (2017) 307-318.

40

[123] H. Wei, J. Ahn, Y.Q. Yu, A. Tymiak, J.R. Engen, G. Chen, Using hydrogen/deuterium exchange mass spectrometry to study conformational changes in granulocyte colony stimulating factor upon PEGylation, J Am Soc Mass Spectrom 23(3) (2012) 498-504. [124] J. Wypych, M. Li, A. Guo, Z. Zhang, T. Martinez, M.J. Allen, S. Fodor, D.N. Kelner, G.C. Flynn, Y.D. Liu, P.V. Bondarenko, M.S. Ricci, T.M. Dillon, A. Balland, Human IgG2 antibodies display disulfide-mediated structural isoforms, J Biol Chem 283(23) (2008) 16194-205. [125] T.M. Dillon, M.S. Ricci, C. Vezina, G.C. Flynn, Y.D. Liu, D.S. Rehder, M. Plant, B. Henkle, Y. Li, S. Deechongkit, B. Varnum, J. Wypych, A. Balland, P.V. Bondarenko, Structural and functional characterization of disulfide isoforms of the human IgG2 subclass, J Biol Chem 283(23) (2008) 16206-15. [126] A. Guo, M. Han, T. Martinez, R.R. Ketchem, S. Novick, C. Jochheim, A. Balland, Electrophoretic evidence for the presence of structural isoforms specific for the IgG2 isotype, Electrophoresis 29(12) (2008) 2550-6. [127] D. Bagal, J.F. Valliere-Douglass, A. Balland, P.D. Schnier, Resolving disulfide structural isoforms of IgG2 monoclonal antibodies by ion mobility mass spectrometry, Anal Chem 82(16) (2010) 6751-5. [128] L.M. Jones, H. Zhang, W. Cui, S. Kumar, J.B. Sperry, J.A. Carroll, M.L. Gross, Complementary MS methods assist conformational characterization of antibodies with altered S-S bonding networks, J Am Soc Mass Spectrom 24(6) (2013) 835-45. [129] A. Zhang, J. Fang, R.Y. Chou, P.V. Bondarenko, Z. Zhang, Conformational difference in human IgG2 disulfide isoforms revealed by hydrogen/deuterium exchange mass spectrometry, Biochemistry 54(10) (2015) 1956-62. [130] Y. Yan, H. Wei, S. Jusuf, S.R. Krystek, Jr., J. Chen, G. Chen, R.T. Ludwig, L. Tao, T.K. Das, Mapping the Binding Interface in a Noncovalent Size Variant of a Monoclonal Antibody Using Native Mass Spectrometry, Hydrogen-Deuterium Exchange Mass Spectrometry, and Computational Analysis, J Pharm Sci 106(11) (2017) 3222-3229. [131] L. Zhang, W. Lilyestrom, C. Li, T. Scherer, R. van Reis, B. Zhang, Revealing a positive charge patch on a recombinant monoclonal antibody by chemical labeling and mass spectrometry, Anal Chem 83(22) (2011) 8501-8. [132] J.C. Geoghegan, R. Fleming, M. Damschroder, S.M. Bishop, H.A. Sathish, R. Esfandiary, Mitigation of reversible self-association and viscosity in a human IgG1 monoclonal antibody by rational, structureguided Fv engineering, mAbs, Taylor & Francis, 2016, pp. 941-950. [133] C.L. Dobson, P.W. Devine, J.J. Phillips, D.R. Higazi, C. Lloyd, B. Popovic, J. Arnold, A. Buchanan, A. Lewis, J. Goodman, Engineering the surface properties of a human monoclonal antibody prevents selfassociation and rapid clearance in vivo, Scientific reports 6 (2016) 38644. [134] T.A. Bowden, K. Baruah, C.H. Coles, D.J. Harvey, X. Yu, B.D. Song, D.I. Stuart, A.R. Aricescu, C.N. Scanlan, E.Y. Jones, M. Crispin, Chemical and structural analysis of an antibody folding intermediate trapped during glycan biosynthesis, J Am Chem Soc 134(42) (2012) 17554-63. [135] Y. Mimura, S. Church, R. Ghirlando, P.R. Ashton, S. Dong, M. Goodall, J. Lund, R. Jefferis, The influence of glycosylation on the thermal stability and effector function expression of human IgG1-Fc: properties of a series of truncated glycoforms, Mol Immunol 37(12-13) (2000) 697-706. [136] A.M. Goetze, Y.D. Liu, Z. Zhang, B. Shah, E. Lee, P.V. Bondarenko, G.C. Flynn, High-mannose glycans on the Fc region of therapeutic IgG antibodies increase serum clearance in humans, Glycobiology 21(7) (2011) 949-59. [137] L. Alessandri, D. Ouellette, A. Acquah, M. Rieser, D. Leblond, M. Saltarelli, C. Radziejewski, T. Fujimori, I. Correia, Increased serum clearance of oligomannose species present on a human IgG1 molecule, MAbs 4(4) (2012) 509-20. [138] T.S. Raju, Terminal sugars of Fc glycans influence antibody effector functions of IgGs, Curr Opin Immunol 20(4) (2008) 471-8.

41

[139] R. Jefferis, Recombinant antibody therapeutics: the impact of glycosylation on mechanisms of action, Trends Pharmacol Sci 30(7) (2009) 356-62. [140] I. Quast, J.D. Lunemann, Fc glycan-modulated immunoglobulin G effector functions, J Clin Immunol 34 Suppl 1 (2014) S51-5. [141] T.P. Patel, R.B. Parekh, B.J. Moellering, C.P. Prior, Different culture methods lead to differences in glycosylation of a murine IgG monoclonal antibody, Biochem J 285 ( Pt 3) (1992) 839-45. [142] M.R. Lifely, C. Hale, S. Boyce, M.J. Keen, J. Phillips, Glycosylation and biological activity of CAMPATH1H expressed in different cell lines and grown under different culture conditions, Glycobiology 5(8) (1995) 813-22. [143] N. Jenkins, R.B. Parekh, D.C. James, Getting the glycosylation right: implications for the biotechnology industry, Nat Biotechnol 14(8) (1996) 975-81. [144] P.H. van Berkel, J. Gerritsen, G. Perdok, J. Valbjorn, T. Vink, J.G. van de Winkel, P.W. Parren, N-linked glycosylation is an important parameter for optimal selection of cell lines producing biopharmaceutical human IgG, Biotechnol Prog 25(1) (2009) 244-51. [145] H.H. Shi, C.T. Goudar, Recent advances in the understanding of biological implications and modulation methodologies of monoclonal antibody N-linked high mannose glycans, Biotechnol Bioeng 111(10) (2014) 1907-19. [146] D. Houde, J. Arndt, W. Domeier, S. Berkowitz, J.R. Engen, Characterization of IgG1 Conformation and Conformational Dynamics by Hydrogen/Deuterium Exchange Mass Spectrometry, Anal Chem 81(14) (2009) 5966. [147] S. Radaev, S. Motyka, W.H. Fridman, C. Sautes-Fridman, P.D. Sun, The structure of a human type III Fcgamma receptor in complex with Fc, J Biol Chem 276(19) (2001) 16469-77. [148] D. Houde, Y. Peng, S.A. Berkowitz, J.R. Engen, Post-translational modifications differentially affect IgG1 conformation and receptor binding, Mol Cell Proteomics 9(8) (2010) 1716-28. [149] P.F. Jensen, V. Larraillet, T. Schlothauer, H. Kettenberger, M. Hilger, K.D. Rand, Investigating the interaction between the neonatal Fc receptor and monoclonal antibody variants by hydrogen/deuterium exchange mass spectrometry, Mol Cell Proteomics 14(1) (2015) 148-61. [150] R.J. Rose, P.H. van Berkel, E.T. van den Bremer, A.F. Labrijn, T. Vink, J. Schuurman, A.J. Heck, P.W. Parren, Mutation of Y407 in the CH3 domain dramatically alters glycosylation and structure of human IgG, MAbs 5(2) (2013) 219-28. [151] J. Fang, J. Richardson, Z. Du, Z. Zhang, Effect of Fc-Glycan Structure on the Conformational Stability of IgG Revealed by Hydrogen/Deuterium Exchange and Limited Proteolysis, Biochemistry 55(6) (2016) 860-8. [152] D. Liu, D. Ren, H. Huang, J. Dankberg, R. Rosenfeld, M.J. Cocco, L. Li, D.N. Brems, R.L. Remmele, Jr., Structure and stability changes of human IgG1 Fc as a consequence of methionine oxidation, Biochemistry 47(18) (2008) 5088-100. [153] H. Pan, K. Chen, L. Chu, F. Kinderman, I. Apostol, G. Huang, Methionine oxidation in human IgG2 Fc decreases binding affinities to protein A and FcRn, Protein Sci 18(2) (2009) 424-33. [154] A. Bertolotti-Ciarlet, W. Wang, R. Lownes, P. Pristatsky, Y. Fang, T. McKelvey, Y. Li, Y. Li, J. Drummond, T. Prueksaritanont, J. Vlasak, Impact of methionine oxidation on the binding of human IgG1 to Fc Rn and Fc gamma receptors, Mol Immunol 46(8-9) (2009) 1878-82. [155] W. Wang, J. Vlasak, Y. Li, P. Pristatsky, Y. Fang, T. Pittman, J. Roman, Y. Wang, T. Prueksaritanont, R. Ionescu, Impact of methionine oxidation in human IgG1 Fc on serum half-life of monoclonal antibodies, Mol Immunol 48(6-7) (2011) 860-6. [156] M. Haberger, A.K. Heidenreich, T. Schlothauer, M. Hook, J. Gassner, K. Bomans, M. Yegres, A. Zwick, B. Zimmermann, H. Wegele, L. Bonnington, D. Reusch, P. Bulau, Functional assessment of antibody oxidation by native mass spectrometry, MAbs 7(5) (2015) 891-900.

42

[157] J. Stracke, T. Emrich, P. Rueger, T. Schlothauer, L. Kling, A. Knaupp, H. Hertenberger, A. Wolfert, C. Spick, W. Lau, G. Drabner, U. Reiff, H. Koll, A. Papadimitriou, A novel approach to investigate the effect of methionine oxidation on pharmacokinetic properties of therapeutic antibodies, MAbs 6(5) (2014) 122942. [158] J. Pollastrini, T.M. Dillon, P. Bondarenko, R.Y. Chou, Field flow fractionation for assessing neonatal Fc receptor and Fcgamma receptor binding to monoclonal antibodies in solution, Anal Biochem 414(1) (2011) 88-98. [159] W. Burkitt, P. Domann, G. O'Connor, Conformational changes in oxidatively stressed monoclonal antibodies studied by hydrogen exchange mass spectrometry, Protein Sci 19(4) (2010) 826-35. [160] A. Zhang, P. Hu, P. MacGregor, Y. Xue, H. Fan, P. Suchecki, L. Olszewski, A. Liu, Understanding the conformational impact of chemical modifications on monoclonal antibodies with diverse sequence variation using hydrogen/deuterium exchange mass spectrometry and structural modeling, Anal Chem 86(7) (2014) 3468-75. [161] L. Bonnington, I. Lindner, U. Gilles, T. Kailich, D. Reusch, P. Bulau, Application of Hydrogen/Deuterium Exchange-Mass Spectrometry to Biopharmaceutical Development Requirements: Improved Sensitivity to Detection of Conformational Changes, Anal Chem 89(16) (2017) 8233-8237. [162] L. Tang, S. Sundaram, J. Zhang, P. Carlson, A. Matathia, B. Parekh, Q. Zhou, M.C. Hsieh, Conformational characterization of the charge variants of a human IgG1 monoclonal antibody using H/D exchange mass spectrometry, MAbs 5(1) (2013) 114-25. [163] H. Ghasriani, D.J. Hodgson, R.G. Brinson, I. McEwen, L.F. Buhse, S. Kozlowski, J.P. Marino, Y. Aubin, D.A. Keire, Precision and robustness of 2D-NMR for structure assessment of filgrastim biosimilars, Nat Biotechnol 34(2) (2016) 139-41. [164] B. Japelj, G. Ilc, J. Marusic, J. Sencar, D. Kuzman, J. Plavec, Biosimilar structural comparability assessment by NMR: from small proteins to monoclonal antibodies, Sci Rep 6 (2016) 32201. [165] L.W. Arbogast, R.G. Brinson, T. Formolo, J.T. Hoopes, J.P. Marino, 2D (1)H(N), (15)N Correlated NMR Methods at Natural Abundance for Obtaining Structural Maps and Statistical Comparability of Monoclonal Antibodies, Pharm Res 33(2) (2016) 462-75. [166] R. Ramanathan, M.L. Gross, W.L. Zielinski, T.P. Layloff, Monitoring recombinant protein drugs: a study of insulin by H/D exchange and electrospray ionization mass spectrometry, Anal Chem 69(24) (1997) 5142-5. [167] S. Nakazawa, N. Hashii, A. Harazono, N. Kawasaki, Analysis of oligomeric stability of insulin analogs using hydrogen/deuterium exchange mass spectrometry, Anal Biochem 420(1) (2012) 61-7. [168] S. Nakazawa, J. Ahn, N. Hashii, K. Hirose, N. Kawasaki, Analysis of the local dynamics of human insulin and a rapid-acting insulin analog by hydrogen/deuterium exchange mass spectrometry, Biochim Biophys Acta 1834(6) (2013) 1210-4. [169] D. Houde, S.A. Berkowitz, J.R. Engen, The utility of hydrogen/deuterium exchange mass spectrometry in biopharmaceutical comparability studies, J Pharm Sci 100(6) (2011) 2071-86. [170] I.A. Kaltashov, C.E. Bobst, R.R. Abzalimov, S.A. Berkowitz, D. Houde, Conformation and dynamics of biopharmaceuticals: transition of mass spectrometry-based tools from academe to industry, J Am Soc Mass Spectrom 21(3) (2010) 323-37. [171] D. Houde, S.A. Berkowitz, Conformational comparability of factor IX-Fc fusion protein, factor IX, and purified Fc fragment in the absence and presence of calcium, J Pharm Sci 101(5) (2012) 1688-700. [172] J. Visser, I. Feuerstein, T. Stangler, T. Schmiederer, C. Fritsch, M. Schiestl, Physicochemical and functional comparability between the proposed biosimilar rituximab GP2013 and originator rituximab, BioDrugs 27(5) (2013) 495-507. [173] J. Fang, C. Doneanu, W.R. Alley, Jr., Y.Q. Yu, A. Beck, W. Chen, Advanced assessment of the physicochemical characteristics of Remicade(R) and Inflectra(R) by sensitive LC/MS techniques, MAbs 8(6) (2016) 1021-34. 43

[174] J. Pan, S. Zhang, A. Chou, D.B. Hardie, C.H. Borchers, Fast Comparative Structural Characterization of Intact Therapeutic Antibodies Using Hydrogen-Deuterium Exchange and Electron Transfer Dissociation, Anal Chem 87(12) (2015) 5884-90. [175] M. Bucciantini, E. Giannoni, F. Chiti, F. Baroni, L. Formigli, J. Zurdo, N. Taddei, G. Ramponi, C.M. Dobson, M. Stefani, Inherent toxicity of aggregates implies a common mechanism for protein misfolding diseases, Nature 416(6880) (2002) 507-11. [176] A.S. Rosenberg, Effects of protein aggregates: an immunologic perspective, AAPS J 8(3) (2006) E5017. [177] W. Wang, Protein aggregation and its inhibition in biopharmaceutics, Int J Pharm 289(1-2) (2005) 130. [178] H. Liu, C. Chumsae, G. Gaza-Bulseco, E.R. Goedken, Domain-level stability of an antibody monitored by reduction, differential alkylation, and mass spectrometry analysis, Anal Biochem 400(2) (2010) 244-50. [179] Y. Li, T.D. Williams, E.M. Topp, Effects of excipients on protein conformation in lyophilized solids by hydrogen/deuterium exchange mass spectrometry, Pharm Res 25(2) (2008) 259-67. [180] B.S. Moorthy, S.G. Schultz, S.G. Kim, E.M. Topp, Predicting protein aggregation during storage in lyophilized solids using solid state amide hydrogen/deuterium exchange with mass spectrometric analysis (ssHDX-MS), Mol Pharm 11(6) (2014) 1869-79. [181] B.S. Moorthy, I.E. Zarraga, L. Kumar, B.T. Walters, P. Goldbach, E.M. Topp, A. Allmendinger, SolidState Hydrogen-Deuterium Exchange Mass Spectrometry: Correlation of Deuterium Uptake and LongTerm Stability of Lyophilized Monoclonal Antibody Formulations, Mol Pharm 15(1) (2018) 1-11. [182] E.M. Moussa, N.E. Wilson, Q.T. Zhou, S.K. Singh, S. Nema, E.M. Topp, Effects of Drying Process on an IgG1 Monoclonal Antibody Using Solid-State Hydrogen Deuterium Exchange with Mass Spectrometric Analysis (ssHDX-MS), Pharm Res 35(1) (2018) 12. [183] G. Wang, P.V. Bondarenko, I.A. Kaltashov, Multi-step conformational transitions in heat-treated protein therapeutics can be monitored in real time with temperature-controlled electrospray ionization mass spectrometry, Analyst DOI:10.1039/C7AN01655G (2018). [184] R.E. Iacob, G.M. Bou-Assaf, L. Makowski, J.R. Engen, S.A. Berkowitz, D. Houde, Investigating monoclonal antibody aggregation using a combination of H/DX-MS and other biophysical measurements, J Pharm Sci 102(12) (2013) 4315-29. [185] R. Majumdar, P. Manikwar, J.M. Hickey, H.S. Samra, H.A. Sathish, S.M. Bishop, C.R. Middaugh, D.B. Volkin, D.D. Weis, Effects of salts from the Hofmeister series on the conformational stability, aggregation propensity, and local flexibility of an IgG1 monoclonal antibody, Biochemistry 52(19) (2013) 3376-89. [186] P. Manikwar, R. Majumdar, J.M. Hickey, S.V. Thakkar, H.S. Samra, H.A. Sathish, S.M. Bishop, C.R. Middaugh, D.D. Weis, D.B. Volkin, Correlating excipient effects on conformational and storage stability of an IgG1 monoclonal antibody with local dynamics as measured by hydrogen/deuterium-exchange mass spectrometry, J Pharm Sci 102(7) (2013) 2136-51. [187] R. Majumdar, R. Esfandiary, S.M. Bishop, H.S. Samra, C.R. Middaugh, D.B. Volkin, D.D. Weis, Correlations between changes in conformational dynamics and physical stability in a mutant IgG1 mAb engineered for extended serum half-life, MAbs 7(1) (2015) 84-95. [188] R.Y. Huang, R.E. Iacob, S.R. Krystek, M. Jin, H. Wei, L. Tao, T.K. Das, A.A. Tymiak, J.R. Engen, G. Chen, Characterization of Aggregation Propensity of a Human Fc-Fusion Protein Therapeutic by Hydrogen/Deuterium Exchange Mass Spectrometry, J Am Soc Mass Spectrom 28(5) (2017) 795-802. [189] G. Deperalta, M. Alvarez, C. Bechtel, K. Dong, R. McDonald, V. Ling, Structural analysis of a therapeutic monoclonal antibody dimer by hydroxyl radical footprinting, MAbs 5(1) (2013) 86-101. [190] M. Eisenstein, Something new under the skin, Nat Biotechnol 29(2) (2011) 107-9. [191] S.J. Shire, Z. Shahrokh, J. Liu, Challenges in the development of high protein concentration formulations, J Pharm Sci 93(6) (2004) 1390-402.

44

[192] J. Arora, Y. Hu, R. Esfandiary, H.A. Sathish, S.M. Bishop, S.B. Joshi, C.R. Middaugh, D.B. Volkin, D.D. Weis, Charge-mediated Fab-Fc interactions in an IgG1 antibody induce reversible self-association, cluster formation, and elevated viscosity, MAbs 8(8) (2016) 1561-1574. [193] T.J. Kamerzell, R. Esfandiary, S.B. Joshi, C.R. Middaugh, D.B. Volkin, Protein-excipient interactions: mechanisms and biophysical characterization applied to protein formulation development, Adv Drug Deliv Rev 63(13) (2011) 1118-59. [194] R.L. Baldwin, How Hofmeister ion interactions affect protein stability, Biophys J 71(4) (1996) 205663. [195] F. He, C.E. Woods, G.W. Becker, L.O. Narhi, V.I. Razinkov, High-throughput assessment of thermal and colloidal stability parameters for monoclonal antibody formulations, J Pharm Sci 100(12) (2011) 512641. [196] T.J. Gibson, K. McCarty, I.J. McFadyen, E. Cash, P. Dalmonte, K.D. Hinds, A.A. Dinerman, J.C. Alvarez, D.B. Volkin, Application of a high-throughput screening procedure with PEG-induced precipitation to compare relative protein solubility during formulation development with IgG1 monoclonal antibodies, J Pharm Sci 100(3) (2011) 1009-21. [197] J. Zhang, D.D. Banks, F. He, M.J. Treuheit, G.W. Becker, Effects of sucrose and benzyl alcohol on GCSF conformational dynamics revealed by hydrogen deuterium exchange mass spectrometry, J Pharm Sci 104(5) (2015) 1592-600. [198] R.T. Toth, B.J. Mills, S.B. Joshi, R. Esfandiary, S.M. Bishop, C.R. Middaugh, D.B. Volkin, D.D. Weis, Empirical Correction for Differences in Chemical Exchange Rates in Hydrogen Exchange-Mass Spectrometry Measurements, Anal Chem 89(17) (2017) 8931-8941. [199] Z. E. Nazari, M. van de Weert, G. Bou-Assaf, D. Houde, A. Weiskopf, K. D. Rand, Rapid Conformational Analysis of Protein Drugs in Formulation by Hydrogen/Deuterium Exchange Mass Spectrometry, J Pharm Sci 105(11) (2016) 3269-3277. [200] S. Rogstad, A. Faustino, A. Ruth, D. Keire, M. Boyne, J. Park, A retrospective evaluation of the use of mass spectrometry in FDA biologics license applications, J Am Soc Mass Spectrom 28(5) (2017) 786-794.

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MS-based conformation analysis of recombinant proteins in design, optimization and development of biopharmaceuticals Devrishi Goswami, Jun Zhang, Pavel V. Bondarenko* and Zhongqi Zhang* Process Development, Amgen, 1 Amgen Center Drive, Thousand Oaks, CA 91320

Highlights

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MS-based conformation analysis is increasingly used in biopharmaceutical development

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Methods include native MS, HDX-MS, covalent labeling, cross-linking, etc.

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Each method is briefly reviewed

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These methods contribute at different stages of biopharmaceutical development

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