Getting to the core of protein pharmaceuticals--Comprehensive structure analysis by mass spectrometry.

European Journal of Pharmaceutics and Biopharmaceutics 93 (2015) 95–109

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European Journal of Pharmaceutics and Biopharmaceutics journal homepage: www.elsevier.com/locate/ejpb

Review Article

Getting to the core of protein pharmaceuticals – Comprehensive structure analysis by mass spectrometry Ulrike Leurs, Ulrik H. Mistarz, Kasper D. Rand ⇑ Department of Pharmacy, University of Copenhagen, Universitetsparken 2, DK-2100 Copenhagen, Denmark

a r t i c l e

i n f o

Article history: Received 2 December 2014 Revised 27 February 2015 Accepted in revised form 2 March 2015 Available online 17 March 2015 Keywords: Pharmaceutical analysis Protein pharmaceuticals Biopharmaceuticals Protein structure analysis Mass spectrometry Hydrogen/deuterium exchange mass spectrometry

a b s t r a c t Protein pharmaceuticals are the fastest growing class of novel therapeutic agents, and have been a major research and development focus in the (bio)pharmaceutical industry. Due to their large size and structural diversity, biopharmaceuticals represent a formidable challenge regarding analysis and characterization compared to traditional small molecule drugs. Any changes to the primary, secondary, tertiary or quaternary structure of a protein can potentially impact its function, efficacy and safety. The analysis and characterization of (structural) protein heterogeneity is therefore of utmost importance. Mass spectrometry has evolved as a powerful tool for the characterization of both primary and higher order structures of protein pharmaceuticals. Furthermore, the chemical and physical stability of protein drugs, as well as their pharmacokinetics are nowadays routinely determined by mass spectrometry. Here we review current techniques in primary, secondary and tertiary structure analysis of proteins by mass spectrometry. An overview of established top-down and bottom-up protein analyses will be given, and in particular the use of advanced technologies such as hydrogen/deuterium exchange mass spectrometry (HDX-MS) for higher-order structure analysis will be discussed. Modification and degradation pathways of protein drugs and their detection by mass spectrometry will be described, as well as the growing use of mass spectrometry to assist protein design and biopharmaceutical development. Ó 2015 Elsevier B.V. All rights reserved.

1. Introduction The worldwide annual revenue of biopharmaceuticals is more than $165 billion. With 588 biosimilars and 434 biobetters in development, this number is likely to increase over the coming years [1]. Protein pharmaceuticals are large molecules containing hundreds of amino acids and having molecular masses from tens to hundreds of kDa. Protein pharmaceuticals thus represent a different class of drug compounds to be analyzed and characterized at the molecular level compared to small molecule drugs [2]. Any modification of either the primary, secondary, tertiary or quaternary structure of a protein pharmaceutical can in principle impact its function, efficacy and safety profile [3,4]. It is therefore important to be able to detect and characterize these modifications. Since the development of soft ionization techniques in the 1990s [5–9], mass spectrometry (MS) has developed into a powerful technique to characterize both primary and higher order structures of protein pharmaceuticals [10,11]. These developments of MS have recently been the subject of excellent specialized reviews (ion mobility spectroscopy [12], HDX-MS [13–17], higher-order ⇑ Corresponding author. E-mail address: [email protected] (K.D. Rand). http://dx.doi.org/10.1016/j.ejpb.2015.03.012 0939-6411/Ó 2015 Elsevier B.V. All rights reserved.

structure analysis by MS [18,19]). However, as the analysis of chemical and physical stability and comparability of protein drugs, both in vitro and in vivo, increasingly relies on an expanding number of MS based workflows, a broader and more comprehensive understanding of the application of MS in biopharmaceutical development is needed by the pharmaceutical scientist. In this review, we seek to provide a broad introduction to the use of both established MS workflows and emerging MS technologies for indepth qualitative analysis of proteins from the perspective of pharmaceutical research. 1.1. Mass spectrometry of peptides and proteins With the advent of soft ionization, the analysis of macromolecules such as peptides and proteins by mass spectrometry became feasible. Soft ionization maintains a low internal energy in the analyte throughout the ionization process, ensuring that the backbone polypeptide structure is left intact. The most commonly used soft ionization techniques for protein mass spectrometry are matrix-assisted laser desorption/ionization (MALDI) [7] and electrospray ionization (ESI) [8]; their principle mechanisms are depicted in Fig. 1. In MALDI, the analyte is co-crystallized with a matrix that allows absorption of a certain wavelength. Upon

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U. Leurs et al. / European Journal of Pharmaceutics and Biopharmaceutics 93 (2015) 95–109

Fig. 1. Principles of MALDI and ESI ionization, as well as peptide fragmentation by tandem mass spectrometry leading to the formation of a-, b-, c-, x-, y- and z-ions, as defined by Roepstorff-Fohlmann [22].

irradiation of the sample with a laser under high vacuum, the matrix material absorbs the energy and ablates. Proton transfer reactions in the hot plume are thought to lead to ionization of sample molecules but the exact mechanism of ionization in MALDI remains to be elucidated [20]. In ESI, ions are produced by dispersing the sample solution at the tip of a capillary into a fine aerosol, due to a large potential difference placed between the positive capillary and a negative anode inside the ionization chamber. Desolvation of the microdroplets in the aerosol is achieved by a flow of counter-current heating gas, resulting in a series of coulombic fissions into droplets of decreasing size and the eventual emergence of desolvated analyte ions. However, as is the case for MALDI, the exact mechanism of ESI remains to be elucidated [21]. Once ionized, the mass spectrometer records the m/z value of resulting peptide or protein ions (MS analysis). Tandem mass spectrometry, or MS/MS, can provide additional information about structure and composition of peptides and proteins. In MS/MS, initial MS analysis of the precursor ion is followed by gas-phase activation and subsequent MS analysis of the resulting dissociated ‘‘product’’ ions. Common types of ion activation for peptides and proteins include collision-induced dissociation (CID) [23], electron transfer dissociation (ETD) [24] or electron capture dissociation (ECD) [25]. CID is accomplished by colliding the gas-phase analyte ions with neutral gas atoms (e.g. He, N2, Ar) [26]. A disadvantage of this approach is that labile posttranslational modifications such as

phosphorylation and glycosylation are easily lost because the backbone amide bonds in proteins and peptides require higher collision energies to break, compared to e.g. glycosidic bonds [27]. However, use of either ETD or ECD can circumvent these problems. For ETD, an electron is transferred from a reagent radical anion (e.g. anthracene) to the analyte ion [24], while in the case of ECD a free low energy electron is captured directly by the analyte ion [25]. The prompt nature of radical-induced fragmentation via ECD or ETD prevents the loss of posttranslational modifications enabling localization of PTMs by mass analysis of product ions through a nonergodic process [28,29]. Depending on the type of fragmentation, different kinds of ions are generated; CID leads to b- and y-ions while ETD and ECD result in the formation of c- and z-ions (Fig. 1).

2. Characterization of protein pharmaceuticals – analysis of covalent (primary) structure Primary structure analysis aims to determine protein purity, molecular mass, amino acid sequence and posttranslational modifications, often as a function of expression, purification, formulation, distribution, handling or storage. Fig. 2 provides an overview of general workflows for primary structure analysis of protein pharmaceuticals by mass spectrometry. The choice of an adequate MS workflow can vary depending on the nature of the

Fig. 2. Bottom-up- and top-down analysis for the structural characterization of peptides and protein pharmaceuticals by mass spectrometry. In bottom-up approaches, the protein is first digested into peptides, which are then analyzed by either MS (peptide mapping/peptide mass finger printing) or tandem MS (MS/MS). Top-down approaches include intact mass analysis (the inset shows the deconvoluted mass spectrum, displaying the molecular weight of the protein) and gas-phase fragmentation of the protein by tandem MS.


protein, the formulation and modifications of interest. Most workflows can be grouped into ‘‘top-down’’ or ‘‘bottom-up’’ approaches that involve the analysis of the intact protein or peptide fragments of the protein produced by enzymatic digestion, respectively [30]. ‘‘Top-down’’ and ‘‘bottom-up’’ analyses are carried out either using MALDI-MS or LC-ESI-MS setups. MALDI-MS is often the first choice for primary structure analysis due to ease-of-use, robustness, and high tolerance for salts, buffers and detergents, as well as a straightforward interpretation of the obtained spectra. However, MALDI-MS generally generates singly charged ions; hence, large proteins will have m/z values outside the optimal range of commonly employed mass analyzers. Furthermore, gas-phase fragmentation (MS/MS) of singly charged ions is typically less efficient than of ions carrying multiple charges. In contrast to MALDI-MS, ESI-MS analysis of proteins yields multiply charged ion species, requiring charge-state deconvolution. Nevertheless, ESI-MS remains the method of choice for advanced bottom-up analyses, as it can be easily coupled to LC systems for separation and process automation allowing higher-throughput analyses [31]. Bottom-up and top-down are rather complimentary techniques often used in combination for comprehensive protein analysis [32]. For antibodies, a ‘‘middle-down’’ approach is becoming increasingly popular, where an antibody sample is selectively cleaved at its hinge region by the immunoglobulin-degrading enzyme from Streptococcus pyogenes (IdeS), followed by a reduction and alkylation step before the sample is subjected to MS analysis [33]. 2.1. Bottom-up mass spectrometry Bottom-up analysis entails enzymatic digestion of the protein and subsequent analysis of the resulting peptides by mass spectrometry. The bottom-up approach is based on so-called peptide mapping, also known as peptide mass fingerprinting or protein fingerprinting. A pre-requisite for peptide mapping is that each protein produces a unique set of peptides upon enzymatic digestion owing to the distinct proteolytic cleavage patterns dictated by its amino acid sequence and the choice of the protease. To verify the identity of the protein, the experimentally obtained peptide mass list is compared with a theoretical peptide mass list calculated from databases such as Mascot, Sequest or X!Tandem [34]. Sequence coverage ranging from 5 to 70% allows protein identification, but is not always sufficient for a detailed characterization of the protein drug [35]. To increase the sequence coverage, multiple

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proteases combining different proteolytic cleavage patterns can be employed, and reagents assisting protein unfolding can be added. Furthermore, tandem mass spectrometry (MS/MS) can improve the spatial resolution of a peptide map, allowing accurate identification of peptides from the digested protein, and in particular sublocalization of PTMs to individual amino acids in a given peptide [34]. A schematic workflow for bottom-up analysis of a protein pharmaceutical is shown in Fig. 3, and an overview of the most common proteolytic enzymes used for bottom-up analysis and their cleavage sites is given in Table 1. MALDI-ToF is a popular MS technique for peptide mapping because of simple sample preparation and straight-forward data interpretation. Furthermore, MALDI-ToF characterization can provide excellent sensitivity and mass accuracy if properly calibrated. The combination of an intact mass analysis and a peptide mapping experiment makes MALDI-ToF a valuable technique for protein analysis throughout the whole biopharmaceutical development process. An example of a peptide mapping analysis of bovine serum albumin (BSA) by MALDI-ToF after tryptic digestion is shown in Fig. 4. Coupling of chromatographic systems to ESI-MS is widely used for advanced peptide mapping experiments especially within the field of proteomics. The chromatographic system allows separation of the proteolytic peptides and thereby increases sequence coverage. Especially for complex mixtures, the chromatographic separation allows sensitive, comprehensive and high-throughput identification of protein components [36]. For biopharmaceutical development, LC-ESI-MS is also useful to analyze and quantitate proteins in complex biological matrices such as blood, urine and other samples that represent relatively crude mixtures of different components [37]. For example, tryptic peptides of a protein can be quantified by multiple reaction monitoring (MRM) in a targeted Table 1 Cleavage patterns for the most common proteases used for bottom-up mass spectrometry. Enzyme

Cleavage pattern

Trypsin Chymotrypsin Pepsin

C-terminal of Lys and Arg, except when followed by Pro C-terminal of Tyr, Trp, Phe Unspecific (slight preference for C-terminal of Tyr, Trp, Phe, Leu) N-terminal of Asp and Glu C-terminal of Arg C-terminal of Glu (and Asp) C-terminal of Lys N-terminal of Lys

Asp-N Arg-C Glu-C Lys-C Lys-N

Fig. 3. Primary structure characterization of a protein drug by bottom-up analysis and peptide mapping. After denaturation, reduction and alkylation of possible disulfide bonds, the protein is enzymatically digested, e.g. by trypsin, cleaving C-terminally after Arg and Lys residues. Subsequent mass analysis reveals a distinct list of peptide masses, which can be compared to a theoretical peptide map generated in silico from a database to reveal the sequence coverage and protein identity.

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Fig. 4. Bottom-up (peptide mapping) analysis of bovine serum albumin (BSA). The MALDI-ToF mass spectrum was recorded on peptides from a tryptic digest of BSA. Peptide ions detected in the mass spectrum were mapped to different parts of the BSA sequence by comparing the experimental masses to a list of predicted masses from a theoretical peptide map generated in silico based on the sequence of BSA.

analysis. Stable isotope labeled (SIL) analogues can be employed for precise and robust quantitation, either at the intact protein level as SIL-proteins, or after proteolytic cleavage as SIL-proteolytic peptides [38]. Furthermore, LC-ESI-MS analysis is increasingly replacing ligand-binding assays for the quantitation of biopharmaceuticals in biological matrices due to the improved specificity, faster method development with multiplexing capabilities and the ability to detect degradation products and posttranslational modifications [39–41]. 2.2. Top-down mass spectrometry Bottom-up analysis has a proven track-record for primary structure analysis, but the advent of high-resolution mass spectrometry with improved mass range and resolution now also allows direct primary structure analysis without prior proteolysis. Though the advantages of bottom-up approaches regarding i.e. comprehensive sequence coverage are undisputed, top-down approaches are less laborious and time consuming in terms of sample preparation. Further, sample preparation for bottom-up analysis can, if not properly controlled, introduce artificial modifications such as oxidation of Met, deamidation of Asn residues and disulfide bond reshuffling [42]. Intact mass analysis can conveniently provide information about identity, purity and modifications of protein pharmaceuticals because the sequences of recombinant therapeutic proteins are often known. The experimentally obtained molecular weight can then be compared with the theoretical mass to verify protein sequence, and any deviations from the calculated mass will indicate sequence variations, chemical degradation or posttranslational modifications. The most commonly employed instruments for intact mass analysis are ToF, Orbitrap and Fourier transform ion cyclotron resonance (FT-ICR) instruments due to their high resolution, accuracy and wide mass range. Fig. 5A gives an example of an intact mass analysis; shown are the ESI mass spectra of native and oxidized human insulin, displaying two isotopic distributions for the quadruple charged molecule corresponding to the native and oxidized form. Intact mass analysis has also proven capable of distinguishing between innovator protein drugs and their biosimilars in cases of small differences in covalent structure and thus mass [43]. Top-down analysis of intact proteins can be augmented by coupling it with tandem mass spectrometry, enabling both in-depth

analysis of protein modifications and de novo sequencing of the protein. De novo sequencing denotes the determination of peptide and/or protein sequences through tandem MS analysis. Even though modern mass spectrometers have high mass accuracy and resolution, the comparison of experimental and theoretical peptide masses is often not sufficient to reliably identify a peptide/protein sequence without using tandem MS. For top-down MS/MS analysis, the intact protein is fragmented, e.g. by CID, and the fragment ions are analyzed, allowing localization of covalent modifications to specific amino acids. While CID of intact proteins can lead to uneven fragmentation patterns resulting in low sequence coverage and loss of PTMs, both ETD and ECD, as well as MALDI in-source decay (ISD) have been shown to be valuable alternatives in top-down MS/MS analysis that can provide complementary and more informative fragment ion series, improving sequence coverage, as well as characterization and localization of PTMs [44–46]. However, especially for antibodies, the sequence coverage of ETD-mediated top-down analysis remains low, typically not exceeding 20% [44]. Use of ECD on high-resolution FTICR instruments has been shown to increase sequence coverage to 30–35% [45]. An advantage of top-down MS compared to bottom-up approaches is the ability to select individual coexisting protein isoforms for structural analysis based on their distinct m/ z, which is not possible by bottom-up analysis where peptides from different protein isoforms cannot be differentiated. Especially in the case of recombinant therapeutic proteins, different isoforms and deletion mutants are commonly expressed, and need to be thoroughly analyzed and characterized. Since these isoforms are often hard to isolate chromatographically, top-down analysis is the pre-requisite tool for this kind of analysis [47]. Fig. 5B illustrates a top-down analysis of glucagon-like peptide-1 (GLP-1) by ESI-CID-MS/MS; the comprehensive b- and y-ion series provide detailed sequence information and allow sequencing of a large part of the sequence de novo. 2.3. Monitoring chemical degradation and posttranslational modifications Protein pharmaceuticals are subject to a broad range of modifications during recombinant expression, purification, formulation, distribution and storage. As these modifications in principle can

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Fig. 5. Top-down analysis of insulin and GLP-1. (A) ESI mass spectrum of human insulin. The spectrum shows the isotopic distribution of the 4+ ion of insulin [M+4H]4+; the blue trace corresponds to an oxidized form of insulin, while the red trace represents the native species. The m/z difference between the two isotopic distributions is 4, corresponding to a mass difference of 16 Da and hence the addition of an oxygen atom. (B) Top-down analysis of the 5+ ion of glucagon-like peptide 1 (GLP-1) by tandem ESI-MS analysis by CID. The inset shows the intact mass spectrum. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

affect efficacy, bioavailability and immunogenicity, it is important to analyze and control these parameters during biopharmaceutical development and production processes. Table 2 shows an overview of common posttranslational modifications and chemical degradation pathways, as well as their corresponding mass changes. These modifications, corresponding to particular changes in molecular mass, can be identified and localized to specific amino acids in a protein by bottom-up and/or top-down analyses as described earlier. Chemical degradation and modifications of specific relevance for protein drug development include oxidation, deamidation, glycosylation, disulfide scrambling, covalent aggregation [48] and Cterminal lysine processing of antibodies [49,50]. Oxidation can occur upon UV-irradiation, through exposure to visible light in the presence of photosensitizers and oxidizing reagents such as peroxides and other reactive oxygen species. The presence of metal ions in the sample, such as Fe and Cu can catalyze the oxidation of proteins by peroxides. The side chains of Met, Trp, Cys, His, and Tyr residues are potential oxidation sites in proteins, with Met and Trp being the most common sites for oxidation [53–55]. Mono-oxidation causes a mass shift of +15.99 Da, while di-oxidation of Met, Trp and Cys results in a mass change of +31.99 Da [54]. There is no pH range that prevents oxidation; the optimal pH for oxidation varies from amino acid to amino acid. For instance, His residues undergo increased oxidation at neutral and basic pH [56,57], while Trp and Met residues are more readily oxidized below pH 4. Tyr residues are generally more susceptible to oxidation at higher pH [58]. Oxidation of therapeutic proteins is problematic as it can potentially result in severe side effects and loss of therapeutic efficacy. It has e.g. been shown that metal-catalyzed oxidation of recombinant interferon a2b leads to aggregation and drug product immunogenicity [59]. Deamidation is a common chemical degradation for protein pharmaceuticals, occurring particularly on Asn and, at a much slower rate, on Gln residues [3]. Deamidation of Asn under strong acidic conditions occurs through direct hydrolysis and leads to formation of aspartic acid (Asp) which is only 1 Da heavier than Asn [58]. Under neutral and alkaline conditions, a succinimide intermediate is formed through abstraction of NH3, resulting in a mass change of 17.03 Da, followed by hydrolysis to either aspartic acid or iso-aspartic acid [60]. This succinimide intermediate is rather stable under acidic conditions, and can be detected by MS [61]. Though iso-aspartic acid adds a methylene group onto the protein backbone as shown in Fig. 6, both iso- and normal aspartic acids have a mass difference of +1 Da compared to Asn, which makes them undistinguishable based on mass alone. However, isoAsp

Table 2 Overview of selected peptide and protein modifications that can be determined by mass spectrometry [51,52]. Modification

Mass change

Involved amino acids

Hydrolysis/cleavage Homoserine

Various 29.99

Peptide bonds Formed from M by CNBr treatment S, T, Y, D, C, N C, S, T, N Q D, N C K, R, D, Q, N- and C-terminal N, Q C W C-/N-terminal, N, C, H, K, N, Q, R, I, L, D, E, S, T G, D, K, N, P, F, Y, R, C, H, W M, W, C, H, L, K, P, Y N-terminal K, R, N, P M, W, C, Y W K, C, S, T, N-terminal A, K K, R, C, M, N-terminal K, D, E, M Formed from C by iodoacetamide H, F, W C S, T, Y, C S, T, Y, H, D, C, R C C N, S, T, W S, T N, S, T, W N, S, T, K, W, C, R, Y, N-terminal K K, N, Q, R, S, T N, S, T C N-terminal G K K C C-terminal C C C, H C, H, Y

Dehydration Ammonia loss Pyroglutamic acid Succinimide formation Disulfide bond formation Amidation Deamidation Disulfide bond reduction Kynurenine Methylation Hydroxylation Oxidation Formylation Dimethylation Dioxidation Formylkynurenine Acetylation Trimethylation Carbamylation Carboxylation Carboxyamidomethylcysteine Bromination Selenylation Sulfation Phosphorylation Pyridylethylcysteine Cysteinylation Pentoses Deoxyhexoses Hexosamines Glycation Lipoic acid Heptoses Acetylhexosamine Farnesylation Myristoylation Biotinylation Pyridoxal phosphate Palmitoylation Geranylgeranylation S-Glutathione Heme FAD

18.01 17.03 17.03 15.99 2.01 0.98 0.98 2.02 3.99 14.02 15.99 15.99 27.99 28.03 31.99 31.99 42.01 42.05 43.01 43.99 57.02 77.91 79.92 79.96 79.97 105.06 119.00 132.04 146.06 161.07 162.05 188.03 192.06 203.08 204.19 210.20 226.08 231.03 238.23 272.25 305.07 616.18 783.14

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Fig. 6. Deamidation of asparagine (Asn) under strong acidic or neutral to basic conditions. In a strong acidic environment, the amide side chain can be directly hydrolyzed, leading to formation of aspartate (Asp). Under neutral to basic conditions, a cyclic intermediate is formed through abstraction of NH3 ( 17 Da), which subsequently racemizes and/or hydrolyzes to either aspartate (Asp) or iso-aspartate (isoAsp).

and Asp can be discriminated through their side chains by ECDand ETD-MS/MS; isoAsp residues yield the unique fragmentation ions cn + 57 and ztotal-n + 57, with n being the location of the isoAsp residue [62,63]. Further, isoAsp and Asp can be discriminated by their different retention times in RP-HPLC [64]. Deamidations have been identified and characterized in a number of proteins of therapeutic interest, e.g. Lewis et al. reported in 1970 that human growth hormone is degraded upon base treatment and hypothesized that deamidation of Gln and Asn residues could be a possible cause of degradation [65]. Another study by Fisher et al. in 1981 investigated the stability of crystalline bovine insulin, and found insulin to be more stable at 20 °C than 5 °C with regard to deamidation and polymerization [66]. A more recent study used peptide mapping by LC–MS to characterize and quantify deamidation in a monoclonal IgG1 antibody [67]. Disulfide bond formation is a common and important posttranslational modification of protein pharmaceuticals due to their crucial role in the stabilization of the higher order protein structures by crosslinking the sulfhydryl groups of (conformationally) neighboring Cys residues. The reduction of disulfide bridges and their subsequent recombination with other Cys residues is called disulfide reshuffling or scrambling [42]. This process can impact both tertiary and quaternary protein structure, potentially altering the pharmacodynamics and kinetics of the therapeutic protein or peptide. The detection of disulfide reshuffling is challenging because it does not result in a mass change. Ion mobility mass spectrometry has been employed to analyze disulfide reshuffling as will be discussed later in this review [68]. The number of disulfide bonds in a protein can be determined by mass spectrometry through reducing the protein, resulting in a mass shift of +2.02 Da per reduced disulfide bond. Typical reducing agents include tris(2-carboxyethyl)phosphine (TCEP) or dithiothreitol (DTT) [69]; however, reducing agents are often the cause of significant ion suppression if not removed chromatographically prior to mass analysis. Two recent reports have described the online coupling of an electrochemical cell to mass spectrometric setups, thereby achieving rapid reduction without the addition of reductive reagents and circumventing the problem of ion suppression [70,71]. The localization of disulfide bonds can be determined through ion fragmentation or enzymatic digestion, as disulfide linked proteins yield unique peptides and mass spectrometric

signals. In both approaches, it is important to limit potential disulfide bond reshuffling. The digestion step during a bottom-up MS/ peptide mapping analysis should preferably be performed under acidic conditions, as neutral or alkaline environments can result in rearrangements or reduction of the disulfide linkages. Further, free thiol groups can be alkylated using i.e. iodoacetamide to prevent reformation of disulfide linkages. Glycosylation is one of the most common and biologically significant posttranslational modifications of protein pharmaceuticals. As many therapeutic proteins are based upon serum proteins that are frequently glycosylated in their native state, the correct glycosylation pattern can be of utmost importance for the functionality of protein drugs [72]. Changes in the glycosylation pattern can result in changed solubility, stability and bioavailability, and they can be a determinant of immunogenicity [73]. The analysis of extent and structure of protein glycosylation is therefore of central interest in biopharmaceutical development [74]. Glycans can be classified into N- and O-linked glycans; N-glycans are conjugated to the side-chain amide nitrogen of Asn residues, while O-glycans are conjugated to the hydroxyl groups of Thr or Ser residues [75]. The vast structural diversity and heterogeneity of protein glycosylation make this PTM a formidable analytical challenge. Glycosylation patterns can differ regarding (a) the site of the glycan linkage, (b) the glycan composition, (c) the glycan structure (branched or unbranched) and (d) the glycan length. An efficient and quantitative method for labeling of glycans was developed in 1995 by Bigge et al. utilizing 2-amino benzamide (2-AB) and antranilic acid to label glycans through reductive amination. The 2-AB label is compatible with most downstream enzymatic sample preparation procedures, and labeled glycopeptides and proteins can be analyzed by hydrophilic interaction liquid chromatography (HILIC) or RP-HPLC coupled to fluorescence and/ or mass spectrometric detection [76]. Recently published applications of glycan analysis by 2-AB labeling include the structural characterization of the monoclonal antibody Nimotuzumab that is currently undergoing clinical trials [77]. In recent years, mass spectrometry has become a favored method for protein glycan analysis. Information about the individual glycan structures and glycosylation sites can be obtained from peptide mapping; here, the masses of proteolytic peptides of the intact and the deglycosylated species are compared to decipher


glycosylation sites [78]. MS/MS analysis using ETD and ECD leaves most of the glycan structures intact by cleaving only the peptide backbone; hence, they can be employed to localize glycosylation to the single amino acid level [28,79]. As noted earlier, CID fragmentation breaks both glycosidic and peptide bonds, thereby making this technique complimentary to ETD [80]. The combination of both techniques has been shown to be a promising approach for more comprehensive glycan analysis, while CID provides information about glycan composition, and ETD allows de novo sequencing of peptides and the localization of glycosylation sites [81]. Fig. 7 shows an example of the identification of a glycosylation site of an antibody by intact mass analysis, as well as through peptide mapping by LC–MS/MS [82]. Glycoform analysis is of particular importance in protein drug development when biopharmaceutical innovator products are compared to their biosimilar counterparts [83]. A glycoform of a protein is a protein isoform containing a certain type of glycan, or a distinct combination of glycans. Differences in the glycosylation of a biopharmaceutical can have an important impact on both the safety and efficacy of the biopharmaceutical. The composition of a given glycosylation can be determined through the sequential release of glycans: By comparing the mass of the glycosylated species to its partially and fully deglycosylated counterparts, conclusions regarding the composition and sequence of a glycosylation can be drawn. The glycans can be released by means of enzymatic and chemical reactions: The most commonly used enzymes are PNGase F, sialidase and different endoglycosidases. Glycan release

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is often followed by permethylation, because the methylation of the free hydroxyl groups on the glycans increases mass spectrometric sensitivity, providing more informative MS/MS spectra and stabilizing labile glycans [84]. PNGase F cleaves practically all N-linked glycans, except those that contain fucose at the third position of an N-linked GlcNAc glycan [85]. Sialidase releases terminal sialic acid, and O-glycans can be removed chemically by alkaline b-elimination in the presence of sodium borohydride (NaBH4) [86]. Hydrazinolysis releases all unreduced O- and N-linked glycans, while trifluoromethanesulfonic acid hydrolysis results in complete removal of all glycans. Recently, a method releasing both N- and O-glycans linkages through universal proteolysis and permethylation in one step has been introduced [87]. A comprehensive summary of mass spectrometry based approaches for protein glycoform analysis has also been given by Marino et al. [88]. 3. Characterization of protein pharmaceuticals – analysis of higher-order structure The higher-order structure of biopharmaceuticals is a major determinant of therapeutic function, efficacy, bioavailability and safety. It is therefore important to control and analyze higher-order structure during protein drug development. Mass spectrometry has, apart from its ability to analyze protein primary structure, recently emerged as a powerful tool to characterize the higher order structures of protein pharmaceuticals [19,78]. Several mass spectrometric techniques are available for higher-order structure

Fig. 7. Glycoform analysis of a therapeutic antibody by MS. Top-down MS analysis of a reduced glycosylated (A) and a reduced deglycosylated (B) antibody. The heavy chains of the glycosylated antibody (A) are detected with three different glycoforms (G0F, G1F and G2F). (C) Mass analysis and relative abundance of antibody glycopeptides as detected by peptide mapping LC–MS (Hex = Hexose, GlcNAc = N-Acetylhexosamine, dHex = Fucose). Notably, a longer glycopeptide resulting from a missed clevage by trypsin showed G1F in higher abundance than G0F, thus listed in the table are average % values. Figure modified from [82].

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characterization, including hydrogen/deuterium exchange mass spectrometry (HDX-MS), ion mobility mass spectrometry (IM– MS), native mass spectrometry, chemical cross-linking and oxidative foot printing. IM–MS and HDX-MS are the two techniques that have so far proven to be most adaptable to biopharmaceutical development and will thus be described here in more detail (Fig. 8).

3.1. Ion mobility mass spectrometry IM–MS is the separation of desolvated gas-phase ions based on their charge and collisional cross section and their subsequent identification by mass analysis. To this end, the gas-phase protein ions are guided by a weak static electrical field through a drift cell containing a high pressure of inert gas [89,90]. The ions are separated according to their charge and collisional cross section, the latter closely dictated by higher-order structure. Hence, tightly folded proteins will travel faster through the drift tube than their unfolded counterparts [91], and thus IM–MS can separate and quantify conformational states of a protein. Because ion mobility is a fast, sensitive and automatable method, it is a promising tool for obtaining structural information of protein pharmaceuticals during production, formulation and distribution [92]. While transition from solution phase to the gas phase could alter protein higher-order structure, it has been shown convincingly that both protein complexes and individual proteins can retain their overall tertiary structure and native stoichiometry for tens of milliseconds after desolvation, provided ionization is gentle [93–95]. Numerous biopharmaceutical applications using IM–MS have recently been reported, demonstrating the feasibility of detecting structural differences in e.g. large monoclonal antibodies. A report by Debaene et al. utilized IM–MS to monitor the structural heterogeneity and Fab-arm exchange of two humanized IgG4 antibodies [96]. Bagal et al. investigated disulfide bond heterogeneity among IgG2 monoclonal antibodies by IM–MS, showing that distinct gas-phase conformers can be detected [68]. Also lot-to-lot variations in Nglycosylation profiles of the therapeutic antibody Trastuzumab have been successfully determined by LC–IM–MS following proteolysis by trypsin [97]. Furthermore, IM–MS has been employed to separate protein aggregates based on their collisional cross sections [98,99].

3.2. Hydrogen/deuterium exchange mass spectrometry The hydrogen/deuterium exchange (HDX) of proteins can be used to study the molecular interactions and dynamics of proteins in solution, and its coupling to MS has become an increasingly popular choice for studying protein–protein interactions and comparing protein conformational states [100]. HDX-MS is based upon the rapid exchange of backbone amide hydrogens for deuteriums when a protein is diluted into a deuterated buffer. Since deuterium is heavier than hydrogen by 1 Da, the HDX of a protein can be directly measured by MS [101]. The HDX rate of amide hydrogens depends primarily on their hydrogen bonding status and their degree of solvation, hence structured or conformationally rigid regions defined by stable intramolecular hydrogen bonding will exchange at much slower rates than dynamic or fully exposed regions available for hydrogen bonding with solvent. Even small changes to the structural environment or dynamics of a particular amide site, i.e. rearrangements in the protein or interactions with a ligand often perturb local amide hydrogen bonding and thus alter amide HDX rates. Such changes in higher order structure properties can either be detected at the intact protein level (global HDX analysis), or be localized to the level of peptide segments through proteolysis of the labeled protein (local HDX analysis). Local HDX analysis includes proteolysis of the deuterated protein, often through the unspecific protease pepsin that generates a plethora of peptides with overlapping sequence which can in some cases allow resolution of HDX from peptide level to the single amino acid level. Furthermore, it has recently been demonstrated that the resolution of local HDX analysis can also be improved to single amino acid level by incorporating ECD or ETD fragmentation into the HDX-MS workflow [102–106]. ECD has also been shown to allow residue-level resolution in a local HDX analysis without prior LC separation through coupling to a high resolution FTICR mass spectrometer [107]. A major challenge in protein conformational analysis by HDXMS is the back-exchange of deuterium to hydrogen during sample handling and chromatographic separation. Fortunately, back-exchange can be quenched by lowering the pH to 2.3 and the temperature to 0 °C, thereby increasing the half-life for deuterium/ hydrogen exchange to 30–120 min [101]. Hence, rapid chromatographic separation (short LC runs) at 0 °C can significantly minimize the loss of the deuterium label (i.e. reduce back-exchange)

Fig. 8. Characterizing the higher order structure of protein drugs by mass spectrometry. Ion mobility mass spectrometry (IMS) is performed in the gas-phase inside the mass spectrometer in tandem to mass analysis, whereas HDX-MS analysis uses a specialized bottom-up MS workflow to monitor a chemical reaction, hydrogen/deuterium exchange (HDX), that is performed in solution. The gray and red isotopic envelopes represent signals from undeuterated and deuterated proteins, respectively. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)


and allow accurate and reproducible measurement of HDX. However, the resolving power of traditional RP-HPLC is poor at low temperatures; therefore, ultra-high-performance LC (UHPLC) employing particles smaller than 2 lm can be used for chromatographic separation, resulting in superior resolution, sensitivity and separation speed at low temperatures [108,109]. Further, operating the mass spectrometer to perform MS/MS without precursor ion selection in a data-independent data acquisition mode (DIA or MSE), allows the simultaneous detection and identification of a large number of peptides during short LC runs. In MSE, all ions in a certain mass range are fragmented and detected simultaneously, and through aligning the accurate precursor and fragment ion masses to retention time, peptides can be identified [110]. Thereby MSE can often allow higher identification rates during the short LC runs required to minimize back-exchange during HDX-MS than classical data dependent acquisition (DDA) schemes where the mass spectrometer performs MS/MS of sequentially isolated precursor ions [111]. The primary advantage of HDX-MS compared to other biophysical techniques for conformational analysis is that it only requires minute amounts of sample (a few hundred picomoles), is highly tolerant to buffer systems, and allows analysis of dilute samples in solution (>10 lM). Further, HDX-MS is also very sensitive to detect and estimate the degree of difference in conformational dynamics between two protein forms, as well as to pinpoint the location of such differences. For the above reasons, HDX-MS has within recent years seen increasing applicability throughout the discovery and development process of (bio)pharmaceuticals, including the analysis of (a) the interaction between drug and protein target, (b) the conformational properties and stability of protein pharmaceuticals, and (c) the comparability and batch-tobatch variations between related protein pharmaceuticals (Fig. 9). Also, the introduction of automated HDX-MS workflows, for instance through a LEAP Technologies Twin HTS PAL liquid handling robot, has allowed higher throughput and more reproducible sample handling which in turn has expanded the applicability of HDX-MS in pharmaceutical industry [112].

3.2.1. Ligand interactions and epitope mapping Traditional structure elucidation techniques, such as X-ray crystallography and NMR, offer single atomic resolution. However, certain protein states, and proteins larger than 40 kDa often resist

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crystal formation and can hence not be analyzed by those techniques. Further, dynamic changes in the protein structure, i.e. through ligand binding or as an intrinsic property of the protein, are not readily detected. Solution-phase HDX-MS may detect and characterize such changes, and has therefore within recent years become a popular technique to study the effects of ligand binding to proteins. Typical pharmaceutical applications include the mapping of interaction sites of small molecule ligands on protein targets, the mapping of binding epitopes of potential antibody pharmaceuticals on protein antigens, as well as the study of changes in conformation or dynamics upon ligand–receptor interactions [100,113–115]. Fig. 10A illustrates the common HDX-MS workflow for analysis of protein–ligand interactions. The mapping of antigen epitopes on antibodies is important with regard to the advance of monoclonal antibodies as pharmaceuticals, for both scientific and regulatory, but also intellectual property concerns. The applicability of HDX-MS for epitope mapping has been demonstrated by various groups [116–118], and comparison to X-ray crystallographic analysis has confirmed the reliability of this technique [119]. For the use of HDX-MS to map protein–ligand interactions, it is important to note that changes in HDX detected upon ligand interaction can be due to both direct (intermolecular) and indirect (intramolecular) changes to the stability (dynamics) of amide hydrogen bonds in the target protein. Thus HDX-MS does not merely reveal the static ligand binding site, but rather the dynamic ‘‘perturbation’’ site of a given ligand on the target protein, providing a full solution-phase view of all backbone amide sites perturbed directly or indirectly upon ligand interaction [100,114,120]. Information on the perturbation site of a drug can be highly valuable for a better understanding of drug-target engagement and reveal new opportunities for drug development.

3.2.2. Quality control and comparability analysis The functionality of a protein drug is intimately dependent on conformation. Protein conformation can be altered through e.g. sequence variations, as well as chemical and posttranslational modifications arising from recombinant protein expression, purification, formulation or storage. The continuous quality control of biopharmaceuticals during development is hence essential to ensure safety and efficacy. HDX-MS has analytical features that make the method an attractive option for protein quality control. In comparison with other techniques, sample consumption is

Fig. 9. Applications of HDX-MS in biopharmaceutical drug development. Left panel: Mapping of protein–drug and protein–protein interaction sites and related conformational changes. Middle panel: Monitoring the conformational response of protein pharmaceuticals to formulation excipients, buffer conditions and long-term storage. Right panel: Comparing the conformational properties of a protein drug to a biosimilar product or across different production batches. The asterisks indicate the location of changes in protein conformation or sites perturbed upon inter-molecular interaction.

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Fig. 10. Continuous labeling HDX-MS workflow for (A) mapping the conformational response of a protein target to binding of a ligand/drug candidate and (B) mapping the conformational differences between two states of a therapeutic protein. After dilution into a deuterated buffer, the HDX reaction is quenched after different time points, and the labeled protein sample is enzymatically digested. The proteolytic fragments are analyzed by LC–MS and regions of the protein involved in conformational changes or ligand binding can be identified based on their differential deuterium uptake.

minimal in HDX-MS, requiring only pico- to nanomoles of a protein sample, and there are no limitations regarding the size or three-dimensional structure of the protein; for instance full length antibody/antigen complexes larger than 250 kDa are analyzed routinely. Furthermore, as HDX-MS is sensitive to small perturbations to the conformational ensemble of a protein in solution, a comprehensive solution-phase view of differences in conformation can be provided. Fig. 10B illustrates the common HDXMS workflow for analysis of conformational differences between two protein states (resulting from variation in sequence, production, formulation, storage, etc.). Current limitations of HDX-MS analysis for quality control of protein pharmaceuticals include the time required for data analysis and the assessment of significance which have so far complicated the implementation of high-throughput workflows. Several modes of data display and analysis aiming to simplify the assessment of differences have been described in the literature [121,122]. The specific utility of HDX-MS for comparability studies of protein pharmaceuticals was recently illustrated by Houde et al. [121]. Other examples of quality control of biopharmaceuticals include the study of interferon-b-1a [123], an IgG1 monoclonal antibody [124] and the effects of PEGylation on the conformation of granulocyte colony stimulating factor [122]. HDX-MS has also been employed to detect changes in antibody structure upon

oxidative stress, revealing several structural effects related to methionine oxidation [125]. Apart from quality control during development, improved analysis of biosimilars to show conformational similarity to the innovator product and thus lower the need for additional clinical trials, represents an exciting emerging application area of HDX-MS. This was recently exemplified in a study comparing an innovator mAb to a biosimilar product [126]. 3.2.3. Rational design, glycan engineering and stability optimization Function and stability of protein drugs can be altered through mutation, glycan engineering and chemical modifications. The effects of such modifications on the protein conformation can be detected and characterized by HDX-MS. For example, an HDX-MS study of different mutants of factor VIIa revealed the origin of their intrinsically enhanced activity [127]. The impact of glycosylation on antibody structure and flexibility has also been investigated by HDX-MS, showing that glycosylation plays an important role for the function of antibodies [128,129]. Likewise, conformational differences between charge state variants of IgG have been characterized by HDX-MS [130]. A recent study by Rose et al. investigated the effects of a single point mutation in an IgG1 antibody by HDX-MS and revealed that the mutation not only led to conformational changes within the region of mutagenesis, but also induced allosteric structural effects that possibly led to increased


glycosylation of the antibody [131]. Bobst et al. determined the effects of chemical modification on interferon b-1a in an extensive MS-based analysis. A global HDX-MS analysis showed that the modified protein is more structurally flexible, and local HDX-MS analyses revealed that these large-scale dynamic changes to the protein structure are induced by alkylation of a single cysteine residue [123]. 3.2.4. Formulation design The formulation of a biopharmaceutical will have considerable impact on long-term stability upon storage; e.g. salt concentration, pH and protein concentration affect both the chemical and physical stabilities of a protein drug. By comparing the deuterium uptake kinetics of a protein, information regarding the degree and location of structural changes can be obtained; e.g. two or more different storage conditions and/or buffer compositions can be compared. The effect of different Hofmeister salts (sulfate, chloride and thiocyanate) on the conformational stability of a monoclonal IgG1 antibody has recently been investigated by HDX-MS, showing that thiocyanate destabilized the antibody [132]. Another study investigated the influence of different excipients (sucrose and arginine) on the conformational and storage stability of an IgG1 antibody, indicating that sucrose leads to an overall decrease in protein flexibility and hence stabilizes it against aggregation, while arginine increases flexibility resulting in increased aggregation [133]. A challenge when using HDX-MS to assess the conformational response of a protein to a given formulation is the fact that multiple excipients can interfere with chromatographic separation and mass analysis during the HDX-MS experiment. Furthermore, the chemical rate of exchange (kch) will change as a function of pH, temperature and ionic strength; hence normalization of changes in HDX due solely to changes in the chemical rates as predicted from theory may not be entirely accurate [134,135]. Addressing these challenges in the future will likely increase the use of HDX-MS to guide formulation design of protein pharmaceuticals [16].

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HDX-MS has also been used to study protein pharmaceuticals in amorphous and solid-phase formulations i.e. lyophilized or frozen solution using customized HDX-MS workflows [136,137]. These customized HDX-MS workflows have been employed for demonstration and localization of site-specific effects of common carbohydrate excipients on the conformation of lyophilized proteins [138]. Another example was the study of aggregation mechanisms and structures in the therapeutic mAb Bevacizumab caused by repeated freeze–thaw cycles and thermal stress [139]. 3.2.5. Aggregation analysis During long-term storage and handling, protein pharmaceuticals often form high molecular weight aggregates that can vary considerably in size and shape. Protein aggregation is problematic because it not only diminishes the therapeutic efficacy of the protein drug, but can also lead to adverse reactions such as anaphylactic shock [140]. Protein aggregation often occurs due to stresses during manufacturing, formulation, distribution and storage. Temperature differences, buffer exchange or high protein concentrations can increase the tendency of proteins to aggregate. Hence, protein aggregation has to be monitored closely during the development of therapeutic proteins to ensure their safety and minimize adverse effects. Detection of protein aggregates can be challenging, especially in case of reversible aggregates that may dissociate upon dilution during the analysis. The most commonly used techniques for protein aggregate analysis are size-exclusion chromatography (SEC) [141,142], native gel electrophoresis and dynamic light scattering [143]. SEC has been successfully coupled to (native) ESI-MS detection for the qualitative analysis of protein aggregates [144–146]. HDX-MS has been employed for the analysis of amyloid fibrils characteristic for Alzheimer’s disease and identified regions involved in fibril formation [147]. Finally, Iacob et al. used HDXMS to investigate the conformational impact of aggregation of an antibody [148].

Fig. 11. Analysis of data from an HDX-MS experiment. The mass spectra show the deuterium uptake of a peptic peptide in a continuous labeling HDX-MS experiment after 15 s, 1 min, 10 min, 60 min and 240 min deuteration. The calculated centroid masses are shown in the left panel as blue or red bars. The graph to the right shows the deuterium uptake calculated from these centroid masses plotted against the different time points. The higher deuterium uptake of the peptide in state A compared to state B indicates a disruption of amide hydrogen bonds (increased dynamics) in this region of the protein.

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3.2.6. Data processing and interpretation for HDX-MS analysis The first step in analyzing HDX-MS data is the identification of the proteolytic peptides resulting from digestion of the protein using pepsin. The peptides are normally identified by MS/MS analysis using computational search engines such as Mascot, Sequest and ProteinLynx Global Server (PLGS) [149–151]. In addition, there are HDX-MS software programs including HX-Express [152], DynamX, Hexicon [153], HDX Workbench [154] and HDExaminer that have considerably improved the speed of data analysis. When using protein search engines, it is important to determine the false-positive discovery rate of peptide assignments through a target-decoy database. In such a database, the amino acid sequence of a given protein is reversed, or generated from a random database; the number of matching peptides identified from this database can then be used to estimate the false-positive discovery rate. To further prevent false-positive peptide assignments, rigorous criteria should be set, e.g. a mass accuracy for the intact peptide mass analysis of less than 10 ppm, at least 2 product ions per 10 residues in the precursor ion, an identification rate of 2 out of 3 replicate LC–MS/MS analyses for each peptide and the retention times should not deviate by more than ±1%. The deuterium uptake of each proteolytic peptide can be calculated by subtracting the intensity weighted average mass of the undeuterated peptide from that of the deuterated peptide. An example of processing raw spectral data into deuterium uptake plots is shown in Fig. 11. Multiple replicates should be performed for each sample, to allow the assessment of inter-run deviation and the assignment of significance [113,120]. Also, some data still require particular careful analysis, e.g. the observation of peak broadening which correlates to very slow protein dynamics (so-called EX1 kinetics). The deuterium uptake of peptides displaying this kind of exchange kinetics cannot be determined from centroid mass analysis, but has to be analyzed manually or using specialized software, such as HXExpress [152], and can yield important insights into slower timescale protein dynamics that could be equally important for determining intrinsic functional properties of the studied protein. A comprehensive description of EX1 kinetics and their assessment has been given elsewhere [155].

4. Summary and future perspectives The increasing numbers of protein drugs that enter the pharmaceutical market create a growing need for analytical techniques capable of comprehensive and in-depth qualitative characterization of such large and complex molecules in a reproducible and high-throughput fashion. In recent years, mass spectrometry has become the preeminent tool for primary structure analysis of proteins and a growing number of diverse MS approaches are being implemented into biopharmaceutical development and quality control workflows for higher-order protein structure analysis. Bottom-up and top-down MS strategies are being routinely employed to dissect the vast chemical complexity of large protein pharmaceuticals. The sensitivity and tolerance to protein size of MS-based workflows is being continuously improved, driven by the sustained development of mass spectrometry instrumentation. While a growing body of work has demonstrated the applicability of more advanced MS techniques, such as HDX-MS and IM/MS for higher-order structure characterization during biopharmaceutical development, these approaches have yet to realize their full potential in terms of routine use. Nevertheless, the particular ability of HDX-MS to monitor the conformational response of a biopharmaceutical to (1) molecular interactions (ligand, aggregation), (2) sequence variation or other covalent modification(s) (expression, purification, storage) and (3) external environment

(formulation, storage), while using only minute amounts of sample at low concentration, should continue to expand the applicability of HDX-MS in (bio)pharmaceutical research. In this context we note that recent advances in automated sample preparation and data analysis have already cleared some of the obstacles for more routine use of HDX-MS in drug development research, while others remain to be addressed by future developments. While sample preparation and data acquisition in HDX-MS can become an automated process through the introduction of a sample handling robot, data mining and statistical analysis of large (replicate) data sets remain challenging for fully automated software systems. Though there have been great improvements in software for automated analysis of HDX-MS, efficient data analysis still requires some extent of manual intervention. Similarly, the experimental protocol for HDX-MS (e.g. quench conditions, labeling time) needs to be optimized and changed for every protein system to guarantee optimal results. Hence, in regulatory biopharmaceutical environments, where methods should be robust and easy-to-use, there is a need for further standardization and automation of both experimental methods and subsequent data analysis for probing the conformational integrity of protein pharmaceuticals by HDX-MS. Acknowledgments K.D.R. acknowledges support from The Marie Curie Actions Programme of the E.U. (Grant PCIG09-GA-2011-294214) and the Danish Council for Independent Research | Natural Sciences (Steno Grant 11-104058). References [1] Ronald R. Rader (Ed.), Biosimilars/Biobetters Pipeline Review, 2014. . [2] D.J. Crommelin, G. Storm, R. Verrijk, L. de Leede, W. Jiskoot, W.E. Hennink, Shifting paradigms: biopharmaceuticals versus low molecular weight drugs, Int. J. Pharm. 266 (2003) 3–16. [3] L. Tao, M. Ackerman, W. Wu, P. Liu, R. Russell, Characterization of impurities and degradants in protein therapeutics by mass spectrometry, in: B.N. Pramanik, H.-N. Lee, S.K. Kim, G. Chen (Eds.), Characterization of Impurities and Degradants using Mass Spectrometry, John Wiley & Sons, Inc., 2011. [4] S. Hermeling, D.J. Crommelin, H. Schellekens, W. Jiskoot, Structure– immunogenicity relationships of therapeutic proteins, Pharm. Res. 21 (2004) 897–903. [5] F. Hillenkamp, M. Karas, Mass spectrometry of peptides and proteins by matrix-assisted ultraviolet laser desorption/ionization, Methods Enzymol. 193 (1990) 280–295. [6] F. Hillenkamp, M. Karas, R.C. Beavis, B.T. Chait, Matrix-assisted laser desorption/ionization mass spectrometry of biopolymers, Anal. Chem. 63 (1991) 1193A–1203A. [7] M. Karas, F. Hillenkamp, Laser desorption ionization of proteins with molecular masses exceeding 10,000 daltons, Anal. Chem. 60 (1988) 2299– 2301. [8] J.B. Fenn, M. Mann, C.K. Meng, S.F. Wong, C.M. Whitehouse, Electrospray ionization for mass spectrometry of large biomolecules, Science 246 (1989) 64–71. [9] K. Tanaka, The origin of macromolecule ionization by laser irradiation (Nobel lecture), Angew. Chem. Int. Ed. 42 (2003) 3860–3870. [10] M.L. Gross, G. Chen, B.N. Pramanik, Protein and Peptide Mass Spectrometry in Drug Discovery, Wiley, Hoboken, NJ, 2012. [11] G. Chen, Characterization of Protein Therapeutics using Mass Spectrometry, first ed., Springer, US, 2013. [12] F. Lanucara, S.W. Holman, C.J. Gray, C.E. Eyers, The power of ion mobilitymass spectrometry for structural characterization and the study of conformational dynamics, Nat. Chem. 6 (2014) 281–294. [13] L. Konermann, J. Pan, Y.H. Liu, Hydrogen exchange mass spectrometry for studying protein structure and dynamics, Chem. Soc. Rev. 40 (2011) 1224– 1234. [14] R.E. Iacob, J.R. Engen, Hydrogen exchange mass spectrometry: are we out of the quicksand?, J Am. Soc. Mass Spectrom. 23 (2012) 1003–1010. [15] D.P. Marciano, V. Dharmarajan, P.R. Griffin, HDX-MS guided drug discovery: small molecules and biopharmaceuticals, Curr. Opin. Struct. Biol. 28C (2014) 105–111. [16] R. Majumdar, C.R. Middaugh, D.D. Weis, D.B. Volkin, Hydrogen–deuterium exchange mass spectrometry as an emerging analytical tool for stabilization and formulation development of therapeutic monoclonal antibodies, J. Pharm. Sci. 104 (2014) 327–345.

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Resolution of protein structure by mass spectrometry.

Beyond gene sequencing: analysis of protein structure with mass spectrometry.

Oxidative protein labeling with analysis by mass spectrometry for the study of structure, folding, and dynamics.

Probing the structure of human protein disulfide isomerase by chemical cross-linking combined with mass spectrometry.

Revealing Higher Order Protein Structure Using Mass Spectrometry.

Mass spectrometry coupled experiments and protein structure modeling methods.

Analysis of Electroblotted Proteins by Mass Spectrometry.

deuterium exchange mass spectrometry.

Structure elucidation of DNA-protein crosslinks by using reductive desulfurization and liquid chromatography-tandem mass spectrometry.

Clinical protein mass spectrometry.

Structure of mycobacterial bis-cyclopropane mycolates by mass spectrometry.

Structure Determination of Natural Products by Mass Spectrometry.

Examining the Influence of Phosphorylation on Peptide Ion Structure by Ion Mobility Spectrometry-Mass Spectrometry.

Immunoaffinity enrichment and mass spectrometry analysis of protein methylation.

Affinity purification-mass spectrometry and network analysis to understand protein-protein interactions.

Crux: rapid open source protein tandem mass spectrometry analysis.

refolding analysis by mass spectrometry. Scrambling of disulphide bridges in insulin.

deuterium exchange mass spectrometry.

mass spectrometry.

Detection of hepatitis C virus core protein in serum by atomic force microscopy combined with mass spectrometry.

Removal of SDS from biological protein digests for proteomic analysis by mass spectrometry.

Analysis of peptides of human seminal plasma by mass spectrometry.

Mass spectrometry of protein complexes: from origins to applications.

Mass spectrometry detection of isolevuglandin adduction to specific protein residues.