Characterization of Supramolecular Peptide-Polymer

Home , Quadrupole ion trap, Quadrupole mass analyzer

CHARACTERIZATION OF SUPRAMOLECULAR PEPTIDE-POLYMER

BIOCONJUGATES USING MULTISTAGE TANDEM MASS SPECTROMETRY

A Thesis

Presented to

The Graduate Faculty of the University of Akron

In Partial Fulfillment

of the Requirements for the Degree

Master of Polymer Science

Benqian Wei

May 2019 CHARACTERIZATION OF SUPRAMOLECULAR PEPTIDE-POLYMER

BIOCONJUGATES USING MULTISTAGE TANDEM MASS SPECTROMETRY

Benqian Wei

Thesis

Approved: Accepted:

______Advisor Interim Dean of the College Dr. Chrys Wesdemiotis Dr. Ali Dhinojwala

______Faculty Reader Dean of the Graduate School Dr. Toshikazu Miyoshi Dr. Chand K. Midha

______Department Chair Date Dr. Tianbo Liu

ii ABSTRACT

Mass spectrometry (MS) is an essential analytical tool for the characterization of the structure of biological macromolecules, including protein-protein and protein-ligand complexes. One-dimensional MS separates gas-phase analyte ions based on their mass to charge ratio (m/z); however, to obtain more detailed structural information, tandem MS

(MS/MS), which involves isolation and subsequent fragmentation of a precursor ion, is required.

In this thesis, electrospray ionization multistage tandem mass spectrometry (ESI-

MSn) was employed to examine the non-covalent complexes between poly(styrene sulfonate) (PSS) and poly-L-lysine (PLL). During single-stage ion activation, the PLL peptide chain mainly underwent backbone cleavages without disruption of the non- covalent interaction which could only be broken via sequential application of electron transfer dissociation (ETD) and collisionally activated dissociation (CAD), indicating strong binding interactions between the two polyelectrolyte chains. Such binding properties make PSS a potential “non-covalent (supramolecular) label” for determining the surface accessibility of basic residues on a peptide or protein. To probe this premise, non- covalent complexes of substance P and PSS were characterized by ESI-MSn using different ion activation methods. Both MS2 and MS3 experiments on the substance P + PSS complex resulted in the formation of bn (on CAD) or cn (on ETD) fragments attached non-covalently to the intact PSS chain. All peptide fragments containing the intact PSS chain included

Arg1, Lys3, and Gln5, pointing out that these residues, which are located near the N- terminus, are most likely involved in the noncovalent interaction with PSS. In contrast,

iii Gln6 was excluded from this fragment series, attesting a much weaker interaction with PSS due to lesser accessibility. The strong tendency of PSS to bind peptides noncovalently at sites that can be elucidated by MSn demonstrates a proof-of-concept for the capacity of this approach to unveil higher order structure in proteins.

iv ACKNOWLEDGEMENTS

I would like to first express my deepest gratitude to my advisor, Dr. Chrys

Wesdemiotis for his excellent guidance, valuable suggestions and kindness during the process of my M.S. studies at the University of Akron. His knowledge in chemistry and mass spectrometry directed me towards the right path to achieve my goals during these two years. I would also like to thank him for his help and suggestions regarding my future studies.

I would like to thank my committee member Dr. Toshikazu Miyoshi for taking his time to read my thesis and giving me great advice and feedback. He is such a nice person that I do not hesitate to ask for help whenever I have difficulties. I am also grateful for the

Department of Polymer Science at The University of Akron. Thanks for holding such an excellent program to support me to do research in such a great environment. The faculty and staff here are all so kind to make me feel at home.

I would like to thank all my current and former group members: Dr. Selim

Gerişlioğlu, Kevin Endres, Savannah Snyder, Jason O’Neill, Jialin Mao and Chen Du for their friendship, help and suggestions during these two years. Special thanks go to Dr.

Selim Gerişlioğlu for his assistance and mentoring throughout my studies. He guided me into the world of mass spectrometry. This thesis could not be completed without him.

Lastly, the deepest gratitude goes to my family. I appreciate everything that my parents and family did for me to support my dream to pursue higher level education. I could not finish my M.S. studies without their unconditional love and support.

v TABLE OF CONTENTS

LIST OF FIGURES...... viii

LIST OF SCHEMES ...... xi

CHAPTER

I. INTRODUCTION AND PRE-THESIS REVIEW ...... 1

II. INSTRUMENTAL METHODS AND BACKGROUND ...... 7

2.1. Mass Spectrometry ...... 7

2.1.1. Ionization Techniques ...... 8

2.1.1.1. Electrospray Ionization ...... 8

2.1.1.2. Matrix Assisted Laser Desorption/Ionization ...... 11

2.1.2.1. Quadrupole Mass Analyzer ...... 13

2.1.2.2. Quadrupole Ion Trap Mass Analyzer ...... 14

2.2. Tandem Mass Spectrometry ...... 16

2.2.1. Collisionally Activated Dissociation (CAD) ...... 17

2.2.2. Electron Transfer Dissociation (ETD) ...... 17

III. MATERIALS AND INSTRUMENTATION ...... 20

3.1. Materials ...... 20

3.2. Instrumentation ...... 21

3.2.1. Bruker© HCTultraTM II Quadrupole Ion Trap Mass Spectrometer.... 21

vi IV CHARACTERIZATION OF POLYMER-PEPTIDE SUPRAMOLECULAR BIOCONJUGATES USING MASS SPECTROMETRY ...... 24

4.1. Background ...... 24

4.2. Experimental ...... 26

4.2.1. Sample Preparation ...... 26

4.2.2. Instrumental Conditions ...... 26

4.3. Results and Discussion ...... 27

4.3.1. PLL-PSS Complexes ...... 27

4.3.2. Substance P-PSS Complexes ...... 35

V. CONCLUSIONS ...... 41

REFERENCES ...... 43

APPENDIX ...... 55

vii LIST OF FIGURES Page Figure. 2. 1. Components of a mass spectrometer...... 7

Figure. 2. 2. Diagram of electrospray ionization source (Reproduced from reference 1 with permission)...... 9

Figure. 2. 3. Electrospray ionization mechanisms (Reproduced from reference 83 with permission)...... 10

Figure. 2. 4. Diagram of the principle of MALDI ((Reproduced from reference 1 with permission)...... 12

Figure. 2. 5. Diagram of the principle of a quadrupole mass analyzer (Reproduced from reference 1 with permission)...... 13

Figure. 2. 6. Schematic view of a 3D ion trap, and direction of the x, y and z coordinates.

(Reproduced from reference 1 with permission)...... 16

Figure. 3. 1. Schematic of Bruker© HCTultraTM II quadrupole ion trap mass spectrometer.

(Reproduced from reference 91 with permission)...... 22

Figure. 4. 1. (a) ESI-MS spectrum of the PLL + PSS mixture and (b) zoom-in version of the m/z 500-1000 region. The doubly and triply charged distributions of the PLL-PSS complexes observed are marked on top of the corresponding peaks in red and blue color, respectively. L and S represent the PLL and PSS repeat units, respectively. The peaks at m/z 403.4, 531.4, and 659.6 represent [Ln + H]+ (n = 3-5) ions. (c) A possible structure of

2+ [L6S4 + 2H] (m/z 791.4)...... 27

Figure. 4. 2. Supramolecular PLL-PSS complex between the PLL 8-mer (L8) and the PSS

2+ 4-mer (S4). Possible hydrogen bonding interactions in [L8S4 + 2H] and the major

viii fragment series generated by CAD (bn and yn) and ETD (cn and zn•) are indicated in the structure...... 28

2 2 3+ Figure. 4. 3. (a) MS (CAD) and (b) MS (ETD) spectra of [L8S4 + 3H] (m/z 613.3). A superscripted asterisk (*) indicates fragments that contain the entire PSS chain, bound non- covalently. The sign # denotes consecutive H2O or NH3 losses from bn or bn* fragments.

...... 29

2 2+ 2+ Figure. 4. 4. MS (CAD) spectra of (a) [L5S4 + 2H] (m/z 727.4) and (b) [L6S4 + 2H]

(m/z 791.4.). A superscripted asterisk (*) indicates fragments that contain the entire PSS chain, bound non-covalently. The sign # denotes consecutive H2O or NH3 losses from bn or bn* fragments...... 30

2 2+ 2+ Figure. 4. 5. MS (ETD) spectra of (a) [L5S4 + 2H] (m/z 727.4) and (b) [L6S4 + 2H] (m/z

791.4.), both with zoom-in versions of the m/z regions of major fragment ions. A superscripted asterisk (*) indicates fragments that contain the entire PSS chain, bound non- covalently...... 32

3 + Figure. 4. 6. (a) MS (ETD-CAD) spectra of (a) [L8S4 + 3H - SO3H•] • (m/z 1758.8) and

3+ (b) c7* (m/z 1710.7), both generated by ETD of [L8S4 + 3H] (m/z 613.3), cf. Fig. 4.3b.

Superscripted * or ‡ indicate fragments bound non-covalently to the intact PSS chain (*) or a PSS chain that lost SO3H• (‡), respectively; the latter fragments have the compositions

+ + [bnS4 + H - SO3H•] and [ynS4 + H - SO3H•] . The sign # denotes consecutive H2O or NH3 losses...... 34

Figure. 4. 7. (a) ESI-MS spectrum of the substance P + PSS mixture and (b) zoom-in version of the m/z 960-1400 region. The doubly charged distribution of the substance P-

PSS complexes observed is marked on top of the corresponding peaks in orange color. P

ix symbolizes substance P (undecapeptide with the sequence Arg-Pro-Lys-Pro-Gln-Gln-Phe-

Phe-Gly-Leu-Met or R-P-K-P-Q-Q-F-F-G-L-M) and S the repeat unit of PSS; the end groups of PSS have been omitted for brevity...... 35

2 2 2+ Figure. 4. 8. (a) MS (CAD) and (b) MS (ETD) spectra of [PS4 + 2H] (m/z 1071.9). A superscripted asterisk (*) indicates fragments that contain the entire PSS chain, bound non-

+ covalently. Fragments cn* - 1 and [PS4 + H - SO3H2] are also present in the ETD spectrum.

...... 37

Figure. 4. 9. (a) MS2 (CAD) spectrum of doubly protonated substance P, [P + 2H]2+ (m/z

674.9), and (b) MS2 (ETD) spectrum of triply protonated substance P, [P + 3H]3+ (m/z

450.3). These spectra, which reveal sequence information about Substance P, are different from the corresponding spectra of substance P-PSS complex ions...... 38

3 Figure. 4. 10. MS (ETD-CAD) spectrum of c10* (m/z 2010.6), generated by ETD of [PS4

+ 2H]2+ (m/z 1071.4), cf. Fig. 4.8b. A superscripted asterisk (*) indicates fragments bound non-covalently to the PSS chain. The sign # denotes consecutive H2O or NH3 losses. ... 40

x LIST OF SCHEMES

Page

Scheme. 3. 1. Chemical structure of poly(styrene sulfonate) (PSSn)...... 20

Scheme. 3. 2. Chemical structure of poly-L-lysine (PLL)...... 21

Scheme. 3. 3. Chemical structure and amino acid sequence of substance P...... 21

3+ Scheme. 4. 1. Elimination of a SO3H• (81 Da) radical upon ETD of [L8S4 + 3H] (m/z

613.3)...... 33

xi CHAPTER I

INTRODUCTION AND PRE-THESIS REVIEW

Mass Spectrometry (MS) is a powerful analytical technique to identify unknown chemicals, explore molecular structures and study fundamental principles of chemistry such as ion/ion reactions. It has been proven to be helpful in many fields such as biotechnology, pharmaceuticals, clinical laboratories and environmental studies by enabling the characterization of various kinds of polymers and biomolecules [1,2]. This significance has become more evident after the introduction of soft ionization methods such as matrix assisted laser desorption/ionization (MALDI) [3-5] and electrospray ionization [6]. These soft ionization methods allow the analytes to stay intact during analysis, providing direct molecular information regarding the sample ions. One- dimensional MS provides molecular weight information on gaseous ions by separating them based on their mass-to-charge ratio (m/z). Nevertheless, a detailed structural and conformational information cannot be acquired just from MS analysis. Tandem MS

(MS/MS or MS2) capabilities, which involve isolation and subsequent fragmentation of the molecular precursor ion, are then utilized to reveal these valuable properties of the molecule of interest [1].

MS/MS analysis has been successfully used to characterize the primary sequence of peptides and proteins [7] as well as identify the end groups or backbone connectivity of polymers [8-9]. Many ion activation methods have been developed over the years to accomplish fragmentation of precursor ions including collisionally activated dissociation

(CAD) [10,11], electron capture dissociation (ECD) [12,13], electron transfer dissociation

1 (ETD) [14,15] and different types of photodissociation (PD) [16-18]. The major fragmentation techniques used in this thesis are CAD and ETD, hence only these two methods will be discussed in this chapter and in Chapter II.

Over the decades, CAD has remained the most popular ion activation method even after the development of many other alternative dissociation techniques. This method is carried out by causing energetic collisions between the analyte ions and an inert gas such as nitrogen, helium or argon. This collision process transfers part of the kinetic energy of the ion into internal energy which will eventually lead to fragmentation of the ion of interest

[10]. The main advantage of this workflow is that it is compatible with a wide range of ionization techniques including ESI. With CAD, it is possible to carry out fragmentation of even-electron ions (i.e. ions with no unpaired electrons) which are relatively stable and typically produced by ESI and many other ionization methods [19]. Owing to its high efficiency of energy accumulation and fragmentation, CAD has been the “gold standard” for other ion activation methods as well [7]. However, it still has some disadvantages. The main shortcoming is the stepwise feature of the energy deposition process in CAD, leading to an upper limit of the total energy that can be accumulated throughout the multicollision process. Consequently, cleavages of the weakest bonds often predominate in CAD, and these may involve structurally uninformative neutral losses [7,20]. Another prevailing problem of CAD is the low mass cutoff (LMCO) in ion trap mass spectrometers [21], which refers to the truncation of the low mass range that may result in the missed detection of the lower m/z fragments [7]. Due to these problems, alternative fragmentation techniques such as ECD, ETD and PD have been developed. Among these ion activation methods, ETD has become one of the most popular since its introduction in 2004 [14].

2 One additional disadvantage of CAD is that it may destroy relatively weak non- covalent interactions. Further, for peptides and proteins, higher order structures may be disrupted and labile post-translational modifications (PTMs) may be lost under CAD.

These problems have been addressed by the introduction of electron-based dissociation methods including ECD and ETD. Only ETD will be discussed in this chapter, but these two techniques are strikingly comparable. ETD entails an interaction between multicharged positive analyte ions with negatively charged radical ions, thus causing the transfer of an electron to the analyte ions which leads to charge reduction and fragmentation

[22-24]. Odd-electron radical species are formed through this electron transfer process with little or no vibrational energy redistribution. The energy released through this exoergic process to the charge-reduced peptide or protein radical species results in cleavages at the backbone N-Cα bonds, producing c- and z-type fragments. This is also the major difference compared with CAD which mainly generates b- and y-type fragment ions by cleavages at the more labile amide bonds.

Tandem mass spectrometry (MS2) is employed extensively to investigate the non- covalent interactions in protein-protein and protein-ligand complexes [25-30]. Although the most widely mode of activation used in these studies has been CAD, newer MS2 techniques such as ECD and ETD have proven to be promising alternatives for the elucidation of non-covalent interactions [31,32]. As mentioned above, ETD and ECD dissociate different bonds in the protein chain compared to CAD [14,33], thereby revealing complementary sequence information [34,35], without disrupting relatively weak non- covalent interactions [36,37]. Recent studies have further documented that applying ETD and CAD sequentially (MS3 mode) enhances the extent of fragmentation vis à vis single-

3 stage ETD or CAD [38-40]; this is usually performed by generating the charge-reduced precursor by ETD and subsequently activating further this ion via CAD to increase the fragmentation extent and thus gain more comprehensive structural information [41,42].

The surface of peptides and proteins is the anchoring point for binding other

(bio)molecules to form multimeric complexes [43]. It is therefore crucial to map surface- accessible amino acid residues, as these are the sites developing the non-covalent interactions that ultimately lead to protein-protein or protein-ligand interactions. Mass spectrometry has been the primary analytical tool for the identification of surface- accessible sites labeled by covalent chemical modification or crosslinking [43-54]. In addition to covalent chemical probe methods, other labeling methods such as photoinduced oxidation [55-60], hydrogen−deuterium exchange (HDX) [61-67], and non-covalent labeling [68-70] have also been coupled with tandem mass spectrometry for probing solvent accessibility of proteins in order to reveal more detailed information about the higher-order protein structure. Studies that utilize non-covalent interactions to determine the surface accessibility of proteins are limited. An MS based method using non-covalent interactions to investigate protein structures called selective non-covalent adduct protein probing (SNAPP) was developed by R. Julian et al. [68-70]; however, this method did not examine the surface accessible sites of the proteins.

In chapter II, the fundamental background of mass spectrometry and tandem mass spectrometry is introduced. A mass spectrometer comprises five components: inlet system, ionization source, mass analyzer, ion detector and data system. Two major ionization techniques: electrospray (ESI) and matrix assisted laser desorption/ionization (MALDI) will be introduced in this chapter since they are the most widely used ionization methods 4 for large molecules. The quadrupole ion trap will be the main mass analyzer discussed in this chapter, to explain how mass separation is accomplished. Tandem mass spectrometry fundamentals will also be discussed in detail, as this technique was the main experimental tool for this thesis.

In chapter III, the chemicals used in this thesis are introduced, including all solvents, proteins and polymers. The instrument utilized, a Bruker© HCTultraTM II Quadrupole ion trap mass spectrometer, will also be described in this chapter.

Chapter IV is the main part of this thesis and presents the results and discussion of this research project. The acidic polyelectrolyte poly(styrene sulfonate) (PSS) was evaluated as a “non-covalent label”, to test its suitability as a potential alternative to covalent markers for determining the basic surface-accessible residues on peptides and proteins. First, the non-covalent interactions between PSS and poly-L-lysine (PLL) were examined, since the positively charged lysine side chains are often located on the surface of hydrophilic proteins [43,52]. MS2 experiments on the PLL-PSS complex showed that the non-covalent interaction between these two polyelectrolyte chains is particularly strong and can only be broken via sequential application of ETD and CAD. ETD disrupts a binding site in PLL-PSS, thereby weakening the interaction and enabling breakup of the

PLL-PSS complex on subsequent CAD. The strong binding affinity of PSS for basic sites makes this polyelectrolyte a potential “non-covalent label” for determining the surface accessibility of basic residues on peptides and proteins. To probe this hypothesis, the non- covalent complexes between PSS and the peptide substance P were investigated as a proof- of-concept. As will be shown, multistage tandem mass spectrometry (MSn) involving both

5 ETD and CAD allowed to elucidate the PSS binding location within the non-covalent

(supramolecular) PSS-peptide conjugate.

6 CHAPTER II

INSTRUMENTAL METHODS AND BACKGROUND

2.1. Mass Spectrometry

Mass spectrometry is an analytical technique widely used to identify the composition of molecules and determine the structure of individual substances or complex mixtures. To obtain spectra, the analytes must be converted into gas-phase ions which can be separated by their mass to charge ratio (m/z). A mass spectrometer usually consists of five components: inlet system, ion source, mass analyzer, ion detector and data system

(Figure. 2. 1) [1]. The sample is converted into gas-phase ions in the ionization source where it is introduced by direct injection or chromatography; the ions produced this way are then separated according to their m/z in the mass analyzer. Some mass spectrometers are equipped with multiple mass analyzers for tandem mass spectrometry experiments which are also introduced in this chapter. The separated ions are detected by the ion detector, then converted into electric signals. The data system converts the current to digital information, displaying it as a mass spectrum.

Sample Ionization Mass Ion Data Injection Source Analyzer Detector System

Microchannel Plate Direct Injection EI Quadrupole Electron Multiplier LC(UPLC/HPLC) CI Time of Flight(TOF) GC ESI Quadrupole Ion Trap MALDI ICR Orbitrap

Vaccum Pumps

Figure. 2. 1. Components of a mass spectrometer.

7 Ion creation is a critical step in mass spectrometric analysis. There are several different ionization methods to create gas-phase ions: electron ionization (EI) [71,72], chemical ionization (CI) [73,74], atmospheric pressure chemical ionization (APCI) [75], desorption electrospray ionization (DESI) [76], electrospray ionization (ESI) [6] and matrix-assisted laser desorption/ionization (MALDI) [3-5]. The last two methods are widely used for the characterization of large biomolecules and synthetic polymers since they are “softer” ionization methods that can retain the integrity of analytes during the ionization process. In this study, the ionization method used is ESI; however, both techniques will be discussed in detail.

2.1.1. Ionization Techniques

2.1.1.1. Electrospray Ionization

Electrospray ionization (ESI) is a soft ionization method that was first interfaced with MS in the late 1960s by Dole et al. [6]. The successful application of ESI to the characterization of biological molecules resulted in the award of a Nobel Prize in

Chemistry to John B. Fenn in 2002 [77,78]. ESI has quickly become one of the most popular ionization techniques after its introduction. It was applied to the analysis of biomolecules initially, but its application was later extended to other fields such as synthetic polymers and supramolecules [79-81].

In ESI, the sample is dissolved in a polar solvent and injected through a stainless- steel capillary at a high voltage (3-6kV). The strong electric field leads to charge accumulation at the needle tip, which causes the analyte solution to break apart and form

8 highly charged droplets. The liquid solution aerosolizes as it leaves the capillary at atmospheric pressure. The combinatorial forces of electrostatic repulsion and solvent evaporation induce desolvation of the droplets as they flow into the mass spectrometer, during which ions are shed from the droplets (Figure. 2. 2).

Figure. 2. 2. Diagram of electrospray ionization source (Reproduced from reference 1

with permission).

The mechanism by which charge is transferred from the liquid to the analytes remains a topic in debate. There are two major models to describe this process. The first one is the charge residue mechanism (CRM) which was first introduced by Dole et al.

[6,82,83]. In this model, the droplets undergo shrinkage as the solvents evaporates with the charge remaining unchanged. When the droplets have shrunk to the point that the surface tension is unable to oppose the repelling force from the imposed charges, they deform and explode into smaller droplets. This process repeats until droplets containing a single analyte ion remain. A gas-phase ion can form when the solvent evaporates from this last droplet

(Figure. 2. 3).

Figure. 2. 3. Electrospray ionization mechanisms (Reproduced from reference 83

with permission).

In 1976, another model called ion evaporation model (IEM) was proposed by

Iribarne and Thomson [84], in which the formation of charged droplets is also caused by

Coulombic fission, similar with the previous mechanism. However, this model proposed

10 that with the increase of the electric field strength on the surface of the droplets, direct desorption of single ions out of the droplets would occur (Figure. 2. 3). Both mechanisms are believed to work in concert: the charge residue model dominates for masses higher than

3000 Da while the ion evaporation model is more prevalent for lower masses [85].

Recently, a new model which is suitable for unfolded proteins and disordered, partially hydrophobic chains was introduced [83]. This mechanism describes that these chains will migrate to the surface of the charged droplets and ultimately separate themselves from the droplets. Electrostatic repulsion causes the transfer of charges from the droplets to the extended chains, which are eventually ejected as multiply charged ions

(Figure. 2. 3).

2.1.1.2. Matrix Assisted Laser Desorption/Ionization

MALDI was not implemented in this thesis, but it is a very powerful technique to generate intact gaseous ions and remains one of the most widely used ionization methods since its introduction in 1988 [3], hence it is worth discussing this ionization method in this chapter.

Matrix is a significant component which is usually a small organic molecule that has a strong absorption at the laser wavelength used. The sample preparation process helps to understand the mechanism of MALDI. The analyte is usually mixed with excess amounts of matrix molecules, allowing each analyte molecule to be isolated and surrounded by the matrix. This can prevent the formation of analyte clusters. A small droplet of this mixture is deposited onto a solid plate. As the solvent evaporates, a solid

11 solution of the sample in the matrix is formed, which is then inserted into the vacuum and bombarded with laser light. Due to the strong absorption property of the matrix, the total energy transferred to the sample is controlled. This also makes MALDI a soft ionization technique. The high-energy laser shots heat the sample-matrix mixture and lead to the sublimation and ablation of a portion of the matrix crystals. Intact analyte molecules are vaporized in this heating process and form clusters with the matrix, which will expand into the gas phase. Proton transfer from the photoionized matrix molecules to the analyte molecules is the final step that ionizes the sample [86]. In some cases, a salt solution is also added into the mixture before the deposition of the droplet onto the solid plate to promote the ionization efficiency. For example, Ag+ salts are used for polydienes and polystyrenes; and Na+ salts for O-containing polymers. The principle of MALDI is described in Figure.

2. 4.

Figure. 2. 4. Diagram of the principle of MALDI ((Reproduced from reference 1

with permission).

12 2.1.2. Mass Analyzers

There are many different types of mass analyzers for the separation of gas-phase ions based on their mass-to-charge ratio (m/z). Common analyzers today include the quadrupole, quadrupole ion trap (QIT), time-of-flight (TOF) tube, as well as the orbitrap and Fourier-transform ion cyclotron resonance (FTICR) analyzer. These mass analyzers can be used by themselves or coupled with other mass analyzers. In this section, only the quadrupole and quadrupole ion trap will be discussed since they are the major mass analyzers contained in the instrument utilized in this thesis.

2.1.2.1. Quadrupole Mass Analyzer

The quadrupole mass analyzer was first introduced by German physicists Wolfgang

Paul and Helmut Steinwedel in 1953 [87]. It is one of the most common mass analyzers because of its small size and the capability to be interfaced with other types of mass analyzers.

Figure. 2. 5. Diagram of the principle of a quadrupole mass analyzer (Reproduced

from reference 1 with permission).

13 The quadrupole analyzer is composed of four cylindrical rod electrodes which are placed in parallel to form a path for the transmission of gaseous ions (Figure. 2. 5).

Opposite rods have different polarity in order to generate an oscillating electric field.

Superimopsed radio frequency (RF) and constant direct current (DC) potentials are applied to each set of the rods to create a mass separator. There are two modes for the operation of the quadrupole mass analyzer. When both DC and RF potentials are applied, a quadrupole is acting as a mass filter, only allowing ions within a narrow mass range to be transmitted.

This is useful for the isolation of a specific ion of interest and ejection of all other ions outside the specialized mass range. This mode is employed in tandem MS since it allows for the selection of a precursor ion isolation which is induced to fragment after exiting the quadrupole. Another mode of operation for quadrupoles is the RF-only mode when the DC potential is set to zero. This mode allows all ions within a wide mass range to pass while focusing them as they are transmitted. Ions created by ESI or APCI under atmospheric pressure are forced to drift into the mass analyzer which is held under high vacuum. Ion losses are minimized in this process by using RF-only quadrupoles as focusing lenses. In this thesis, the quadrupole is manly operated in the second mode to focus the ions generated in the ESI source for further analysis in the quadrupole ion trap mass analyzer, which will be introduced in the next section.

2.1.2.2. Quadrupole Ion Trap Mass Analyzer

The quadrupole ion trap (QIT) is a mass analyzer that consists of a ring electrode located between two end caps. The ring electrode is connected to an alternating current

(AC) supply which provides a fundamental RF potential to the trap while the two end caps

14 are grounded (Figure. 2. 6) [88,89]. A three-dimensional quadrupolar field is generated inside the trap by the AC potential to trap ions within a specific m/z range. Ions created in the ionization process are injected into the trap through the small hole in the entrance end cap and move towards to the center of the trap due to the RF field. During the ion accumulation process, ions with the same m/z travel on the same trajectory towards the center where they are stored and trapped. The ions can be ejected through a hole in the exit end cap to reach the detector. This is achieved by increasing the RF voltage applied on the ring electrode which leads to the increase of the amplitude of the ion oscillations, so that the ions be ejected through the hole of the exit end cap.

QIT mass analyzers can be regarded as three-dimensional quadrupoles. One major difference between them is that quadrupoles only allow ions within a specific narrow m/z range to pass through, while QIT mass analyzers are able to store all ions within a wide m/z range and eject them later to the detector based on their m/z values. The pressure inside the analyzer is another difference. Quadrupoles are usually operated under high vacuum; on the other hand, a buffer gas such as helium is introduced into the QIT mass analyzer.

This is due to the fact that the ions trapped in the QIT repel each other and this force accelerates them, destabilizes their trajectories and causes a bad resolution. The buffer gas collides with the ions and cools their kinetic energy. This helps them to keep their own trajectories and reach the detector at the proper ejection fields [1].

In terms of tandem mass spectrometry, a major advantage of QIT analyzers is that they support tandem-in-time rather than tandem-in-space, which is the case for beam instruments. The tandem-in-time is essential for performing multistage tandem mass spectrometry (MSn) experiments, while the tandem-in-space alternative on beam 15

instruments only permits MS2. Moreover, simultaneous accumulation of ions with opposite polarities inside the QIT analyzers makes it possible to carry out various ion/ion reactions which is crucial for electron transfer dissociation (ETD). The latter method will be introduced in the next section.

Figure. 2. 6. Schematic view of a 3D ion trap, and direction of the x, y and z coordinates.

(Reproduced from reference 1 with permission).

2.2. Tandem Mass Spectrometry

Tandem mass spectrometry (MS2) is a two-stage mass analysis technique that isolates a precursor ion which is then fragmented to yield structurally diagnostic product ions and neutral fragments. The fragmentation results can provide sequence, architectural and structural information that cannot be obtained by single stage MS [1,7-18]. In most cases, tandem mass spectrometry experiments utilize two mass analyzers. The first analyzer acts as a mass filter that transmits an isolated precursor ion which undergoes

16 fragmentation in a subsequent collision/reaction cell to form fragment ions that are analyzed by the second one. Fragmentation is achieved by different methods. Most commonly used are collisionally activated dissociation (CAD) and electron transfer dissociation (ETD), which were both performed in this thesis.

2.2.1. Collisionally Activated Dissociation (CAD)

CAD is arguably the most widely used gas-phase fragmentation method for tandem mass spectrometry [7]. Dissociation results from the collision between the ion of interest with an inert gas such as helium, argon, or nitrogen [10].

CAD is typically processed by selecting a precursor ion in the first mass analysis step. For QIT analyzers, this step is performed in the trap by ejecting all the other ions. The isolated ion is induced to undergo energetic collisions with an inert gas. The collisions convert a portion of the kinetic energy of the ion into internal energy which is redistributed along the entire molecule as vibrational energy and eventually causes dissociation of the molecule. The energy deposition in CAD is a stepwise process occurring in multiple collisions, therefore, the total energy that can be accumulated is limited, favoring fragmentation via lower energy pathways that typically lead to cleavages of the most labile bonds.

2.2.2. Electron Transfer Dissociation (ETD)

ETD has become a very promising alternative ion activation method since its introduction in 2004 [14]. It involves ion/ion reactions between a multiply charged analyte

ion with a radical reagent anion. This reaction transfers an electron from the reagent to the ion of interest which ultimately leads to the dissociation of the precursor ion.

[푀 + 푛푋]푛+ + 퐴−• ⟶ [푀 + 푛푋](푛−1)+•∗ ⟶ 푓푟푎푔푚푒푛푡푠

In ETD, the charge state of the precursor ion is particularly significant. A multiply charged cation is required to observe fragmentation as the capture of an electron by a singly charged ion results in a neutral molecule which cannot be detected in a mass spectrometer.

Another important component in ETD is the negatively charged reagent. There is always a competition between electron transfer from the reagent to the precursor ion and proton transfer from the precursor ion to the reagent [14,90]; hence the ability of the reagent to transfer electrons is critical. Many ETD reagents have been studied and ultimately fluoranthene (C16H10) has emerged the best choice in today’s commercial instruments due to its high efficiency for electron transfer.

The internal energy gained in ETD will not be randomized along the whole molecule like in CAD, which prevents that only the weak bonds of the precursor ion dissociate [12,14]. This makes ETD a nonergodic process that preserves the labile bonds in analyte ion. Because of their different mechanisms, CAD and ETD cause different fragmentation pathways and reveal complementary structure information on a molecule.

In addition, the nonergodic feature of ETD makes it useful for investigating proteins and other biomolecules, since it can preserve labile post-translational modifications (PTMs) to a substantially greater extent than CAD [14,90]. This same advantage makes ETD also a crucial tool for the analysis of protein assemblies, since it can cleave the backbone of

18 proteins without disruption of the non-covalent interactions between subunits [7,30]. This characteristic is particularly important for the study discussed in Chapter IV.

CHAPTER III

MATERIALS AND INSTRUMENTATION

3.1. Materials

Solvents, additives, and other reagents used in sample preparation. Both solvents employed for sample preparation (methanol, MeOH and MS grade water) were acquired from Fisher (Fair Lawn, NJ). The buffer (ammonium acetate, NH4OAc) used to protect the non-covalent interactions between the polymer and peptides was also purchased from Fisher (Fair Lawn, NJ).

Polymer reagents. Poly(styrene sulfonate) sodium salt (PSS) carrying the sodium sulfonate group in para position (Mw ≈ 1100 Da) (Scheme. 3. 1) was purchased from

Polymer Standards Service-USA, Inc. (Warwick, RI). This acidic polyelectrolyte was employed as a non-covalent label to determine the strength of the non-covalent interactions with the peptides as well as to determine the basic surface accessible sites of substance P.

Scheme. 3. 1. Chemical structure of poly(styrene sulfonate) (PSSn).

Peptides. Poly-L-lysine hydrobromide with an Mw range from 1000 to 5000 Da

(Scheme. 3. 2) and substance P acetate salt hydrate (≥95% purity) (Scheme. 3. 3) were purchased from Sigma-Aldrich (St. Louis, MO).

20 Scheme. 3. 2. Chemical structure of poly-L-lysine (PLL).

NH2-RPKPQQFFGLM-NH2

Scheme. 3. 3. Chemical structure and amino acid sequence of substance P.

3.2. Instrumentation

3.2.1. Bruker© HCTultraTM II Quadrupole Ion Trap Mass Spectrometer

A Bruker© HCTultraTM II Quadrupole Ion Trap Mass Spectrometer (Figure. 3. 1) equipped with electrospray ionization source (ESI) (Bruker Daltonics, Billerica, MA) was used to carry out the experiments described in chapter IV. The samples were injected into the instrument by direct injection with an external syringe pump (KD Scientific, Holliston,

MA). The sample solution enters the ESI source through a grounded needle encircled by

21 the nebulizer; nitrogen gas streams through the nebulizer and enters the sample solution at the needle tip. The nebulizer gas pressure can be adjusted from 0-80 psi. The heated drying gas (N2) helps solvent evaporation during the ionization process. The ions created by the

ESI source are driven into a glass capillary by the voltage difference between the needle and the entrance of the capillary. The ions pass through the glass capillary which serves as a bridge between atmospheric pressure and vacuum. After exiting the glass capillary, the ions are pushed through the skimmers, octapoles and lenses where they can be focused under vacuum. The voltages of these components are set to a certain value to optimize ion transmission and decrease interference of background noise. The transmitted ions then exit the octopole and enter the holes of the entrance end cap electrode of the QIT mass analyzer which has been discussed in section 2.1.2.2. Only RF potentials are applied to the octapoles which act as focusing lenses in this manner, similar to the RF-only mode quadrupoles discussed in section 2.1.2.1.

Figure. 3. 1. Schematic of Bruker© HCTultraTM II quadrupole ion trap mass spectrometer.

(Reproduced from reference 91 with permission).

The importance of this instrument in this thesis is that it can carry out both CAD and ETD fragmentation. To perform CAD, the precursor ion within a particular m/z range is isolated and undergoes acceleration and collisions with the He gas in the ion trap, leading to dissociation of the precursor ion. The fragment ions are then ejected towards the ion detector after accumulation in the ion trap for enough time. For ETD purpose, gas-phase fluoranthene will react with accelerated electrons in the presence of methane buffer gas in the negative chemical ionization source (nCI). This reaction will generate fluoranthane radical anion reagents which will then enter the ion trap. After enough accumulation of both precursor cations and reagent anions in the trap, ion/ion reactions between these two species occur for a certain amount of time which is set to optimize the dissociation efficiency. The obtained ETD fragment ions are directed to the ion detector. As discussed in section 2.1.2.2 about the QIT mass analyzer, this instrument can be used to operate multistage tandem mass spectrometry (MSn), particularly in this thesis, ETD-CAD. This means that after the first stage ETD, one product ion can be isolated again and undergo a further stage of CAD fragmentation. To perform MSn, all ETD product ions are ejected but one ion of interest is trapped. This precursor ion can then collide with the inert gas and

CAD fragmentation occurs again.

23 CHAPTER IV

CHARACTERIZATION OF POLYMER-PEPTIDE SUPRAMOLECULAR BIOCONJUGATES USING MASS SPECTROMETRY

4.1. Background

Tandem mass spectrometry (MS2) is employed extensively to investigate the non- covalent interactions in protein-protein and protein-ligand complexes [25-30]. Although the most widely mode of activation used in these studies has been collisionally activated dissociation (CAD) [29], newer MS2 techniques such as electron capture dissociation (ECD)

[12] and electron transfer dissociation (ETD) [14] have proven to be promising alternatives for the elucidation of non-covalent interactions[25,31,32]. ETD and ECD dissociate different bonds in the protein chain compared to CAD [14,33]; thereby revealing complementary sequence information [34,35] without disrupting relatively weak non- covalent interactions [36,37]. Recent studies have further documented that applying ETD and CAD sequentially (MS3 mode) enhances the extent of fragmentation vis à vis single- stage ETD or CAD [38-40]; this is usually performed by generating the charge-reduced precursor by ETD and subsequently activating further this ion via CAD to increase the fragmentation extent and thus gain more comprehensive structural information [41,42].

The surface of peptides and proteins is the anchoring point for binding other

(bio)molecules to form multimeric complexes [43]. Consequently, it is crucial to map surface-accessible amino acid residues, as these will participate in the non-covalent interactions that ultimately lead to the multicomponent assemblies. Mass spectrometry has

been the primary analytical tool for the identification of surface-accessible sites labeled by covalent chemical modification or crosslinking [43-54].

In the study presented in this chapter, the acidic polyelectrolyte poly(styrene sulfonate) (PSS) is evaluated as a “non-covalent label”, to test its suitability as a potential alternative to covalent markers for determining the basic surface-accessible residues on peptides and proteins. First, the non-covalent interactions between PSS and poly-l-lysine

(PLL) were examined, since the positively charged lysine side chains are often located on the surface of hydrophilic proteins [43,52]. MS2 experiments on the PLL-PSS complex showed that the non-covalent interaction between these two polyelectrolyte chains is particularly strong and can only be broken via sequential application of ETD and CAD.

ETD disrupts a binding site in PLL-PSS, thereby weakening the interaction and enabling breakup of the PLL-PSS complex on subsequent CAD. The strong binding affinity of PSS for basic sites makes this polyelectrolyte a potential “non-covalent label” for determining the surface accessibility of basic residues on peptides and proteins. To probe this hypothesis, the non-covalent complexes between PSS and the peptide substance P were investigated as a proof-of-concept. As will be shown, multistage tandem mass spectrometry (MSn) involving both ETD and CAD allowed to elucidate the PSS binding location within the non-covalent (supramolecular) PSS-peptide conjugate.

25 4.2. Experimental

4.2.1. Sample Preparation

PLL and PSS solutions were prepared in 50% MeOH at the concentration of 5

μg/mL and were mixed at 1:1 (v/v) ratio to form the non-covalent complexes. For the preparation of PSS-substance P complexes, solutions of substance P and PSS were prepared in 50% MeOH that also contained 15 mM NH4OAc at the concentration of 0.01 mg/mL and 0.02 mg/mL, respectively; the latter solutions were mixed at 1:1 (v/v) ratio.

4.2.2. Instrumental Conditions

All experiments were performed on a Bruker© HCTultraTM II Quadrupole Ion Trap

Mass Spectrometer equipped with an electrospray ionization (ESI) source (Bruker

Daltonics, Billerica, MA). All samples were injected into the ESI source using a syringe pump at a flow rate of 4 μL/min. MS2 and MS3 experiments via CAD or ETD were performed on doubly and triply protonated ions. During CAD experiments, the isolation width was kept at 1.0 Da and the amplitude of the excitation RF field was set between 0.26 and 1.10 (arbitrary units). For ETD, fluoranthene radical anions (reagent ions) were produced in a negative chemical ionization (nCI) source filled with methane buffer gas

(2.0–2.6 bar) and located above the octapole lens that transfers ions from either the ESI or the nCI source to the ion trap. The reagent anion intensity was optimized at the following conditions: reagent ion ICC 100,000; ionization energy 70 eV; emission current 2.0 μA; and reagent remove cutoff m/z 210. After accumulation of both types of species in the ion trap, the ion/ion reaction time was set within 160–170 ms. The reaction time was optimized

26 during the MS2 experiments to maximize the fragment ion abundances for the next fragmentation stage. MS and MSn data were analyzed using Bruker’s Compass

DataAnalysis v.4.0 software.

4.3. Results and Discussion

4.3.1. PLL-PSS Complexes

Figure. 4. 1. (a) ESI-MS spectrum of the PLL + PSS mixture and (b) zoom-in version of the m/z 500-1000 region. The doubly and triply charged distributions of the PLL-PSS

complexes observed are marked on top of the corresponding peaks in red and blue color,

respectively. L and S represent the PLL and PSS repeat units, respectively. The peaks

at m/z 403.4, 531.4, and 659.6 represent [Ln + H]+ (n = 3-5) ions. (c) A possible structure

2+ of [L6S4 + 2H] (m/z 791.4).

The ESI-MS spectrum of solutions containing PLL (H–Ln–OH) and PSS (C4H9–

Sn–H) includes doubly and triply protonated distributions of supramolecular (i.e. non-

2+ covalently bound) PLL-PSS complexes with the composition [LnSm + 2H] and [LnSm +

3H]3+, respectively (cf. Figure. 4. 1). This notation specifies the content in repeat units of

PLL (L, C6H12N2O; 128.09 Da) and PSS (S, C8H8SO3; 184.02 Da); the corresponding end groups (Figure. 4. 2) are omitted for brevity. It is noteworthy that complete –SO3Na → –

SO3H exchange occurs in the PSS side chains when solutions of PSS sodium salt and PLL hydrobromide are mixed to form the supramolecular PLL-PSS bioconjugate.

Figure. 4. 2. Supramolecular PLL-PSS complex between the PLL 8-mer (L8) and the PSS

2+ 4-mer (S4). Possible hydrogen bonding interactions in [L8S4 + 2H] and the major

fragment series generated by CAD (bn and yn) and ETD (cn and zn•) are indicated in the

structure.

2 The MS fragmentation patterns of doubly and triply protonated LnSm complexes in various stoichiometries are very similar, both upon CAD (cf. Figure. 4. 3a vs. Figure. 4.

4) as well as ETD (cf. Figure. 4. 3b vs. Figure. 4. 5). Therefore, only the MS2 and MS3

3+ characteristics of triply protonated L8S4 complex ions, viz. [L8S4 + 3H] , will be discussed.

2 2 3+ Figure. 4. 3. (a) MS (CAD) and (b) MS (ETD) spectra of [L8S4 + 3H] (m/z 613.3).

A superscripted asterisk (*) indicates fragments that contain the entire PSS chain, bound

non-covalently. The sign # denotes consecutive H2O or NH3 losses from bn or bn*

fragments.

3+ CAD of [L8S4 + 3H] causes amide bond cleavages in the PLL backbone, which produce homologous series of bn and yn fragments (Figure. 4. 3a). Consecutive elimination of water or ammonia from the bn fragments also takes place [92,93]. The bn and yn fragment series are observed both with (bn*, yn*) as well as without (bn, yn) the PSS chain. No peaks corresponding to the intact PLL or PSS are present in the spectrum, indicating that the non- covalent interaction between the PLL and PSS polyelectrolyte chains is stronger than the energy required for C-N bond cleavages at the PLL amide bonds.

2 2+ 2+ Figure. 4. 4. MS (CAD) spectra of (a) [L5S4 + 2H] (m/z 727.4) and (b) [L6S4 + 2H]

(m/z 791.4.). A superscripted asterisk (*) indicates fragments that contain the entire PSS

chain, bound non-covalently. The sign # denotes consecutive H2O or NH3 losses from

bn or bn* fragments.

The bn and yn fragments that do not contain the PSS chain are attributed to fragmentation of the doubly protonated PLL-PSS complex by charge separation [92,93] which generates two charged fragments, one bound to PSS non-covalently and the other without the PSS.

Under the low-energy CAD conditions used in this study, multiple collisions occur before fragmentation. The internal energy deposited in each collision is redistributed throughout the whole molecule, until enough internal energy has been accumulated to induce fragmentation within the time spent in the collision cell [94]. This slow increase of

30 internal energy (“slow heating”) promotes dissociations with low energy requirements and mainly cleaves labile bonds [40,95]. The major CAD pathways of PLL-PSS involve cleavages of the relatively weak amide bonds [96] without disruption of the supramolecular complex (Figure. 4. 3a), corroborating the presence of strong non-covalent binding interactions between the PLL and PSS chains.

ETD experiments yield similar results with CAD, as again no dissociation of the supramolecular bioconjugate is observed to form intact PLL or PSS fragments (Figure. 4.

3+ α 3b). ETD of [L8S4 + 3H] primarily induces N–C bond cleavages in the PLL component[12][14], generating cn* and zn* fragments that are bound to the intact PSS chain

(cf. Figure. 4. 3b). A unique fragmentation taking place in ETD is the elimination of a sulfonic acid radical (SO3H•, 81 Da) from the PSS side chain (Figure. 4. 3b and Figure. 4.

5). This dissociation is rationalized in Scheme. 4. 1 via electron transfer to a protonated lysine side chain that is hydrogen-bonded with a PSS side chain. The nascent H• radical created in this step is captured by the sulfonic acid substituent, initiating the loss of SO3H•.

In contrast to CAD, unbound cn or zn fragments have much lower relative abundance in ETD, where the dissociating ions are mainly singly charged; when these ions undergo N–Cα bond scission, the larger fragments (i.e. those containing PSS) more effectively compete for the charge and, thus predominate. Overall, single-stage CAD or

ETD do not break the non-covalent complex, verifying that the PLL and PSS polyelectrolyte chains develop strong intermolecular binding interactions.

2 2+ 2+ Figure. 4. 5. MS (ETD) spectra of (a) [L5S4 + 2H] (m/z 727.4) and (b) [L6S4 + 2H]

(m/z 791.4.), both with zoom-in versions of the m/z regions of major fragment ions. A

superscripted asterisk (*) indicates fragments that contain the entire PSS chain, bound

non-covalently.

+ Two ETD products, viz. the radical ion [L8S4 + 3H - SO3H•] • and the closed-shell ion c7*, were examined by CAD for further information on the structure and stability of the PLL-PSS complex. Sequential ETD-CAD application (MS3) leads to a completely different outcome compared with the MS2 experiments (cf. Figure. 4. 6vs. Figure. 4. 3).

+ CAD of [L8S4 + 3H - SO3H•] • results in C-N bond scissions at the PLL chain, yielding bn‡ and yn‡ fragments attached non-covalently to PSS that lost a -SO3H side chain as well as bn and yn fragments devoid of PSS. It is notable that the most abundant fragment in the

MS3 (ETD-CAD) spectrum is the intact PLL chain (m/z 1043.7); hence, the non-covalent interaction is broken readily after the loss of SO3H•, which reduces the number of binding

32 sites between the two polyelectrolytes. Furthermore, C-C bond scission in the PSS backbone also occurs, with concomitant dehydration, giving rise to the loss of the PSS end group (C4H10, 58 Da) plus water. This latter reaction strongly suggests that the unpaired

+ electron in [L8S4 + 3H - SO3H•] • resides on the PSS segment, from where it can initiate charge-remote homolytic bond cleavages[97] that detach the PSS end group [98].

Scheme. 4. 1. Elimination of a SO3H• (81 Da) radical upon ETD of [L8S4 + 3H]3+ (m/z 613.3).

3 MS (ETD-CAD) experiments on c7* also generate multiple fragment series by amide bond cleavages in the PLL backbone (Figure. 4. 6b), which are observed either with

(bn* and c7yn*) or without (bn and c7yn) the intact PSS chain attached. In addition, the non- covalent PLL-PSS interaction is broken to form the unbound c7 ion; however, the relative abundance of the latter fragment is significantly lower than the relative abundance of PSS-

+ free [L8S4 + 3H - SO3H•] • (cf. c7 in Figure. 4. 6b vs. [L8 + H]+ in Figure. 4. 6a). Evidently, the loss of one SO3H• moiety from the PSS chain weakens markedly more the supramolecular PLL-PSS interaction than the loss of one lysine residue from the PLL chain.

This phenomenon could be a consequence of the larger number of PLL vs. PSS residues in

L8S4, which would allow PSS to find a new PLL side chain for non-covalent interaction if one lysine residue is removed.

3 + Figure. 4. 6. (a) MS (ETD-CAD) spectra of (a) [L8S4 + 3H - SO3H•] • (m/z 1758.8)

3+ and (b) c7* (m/z 1710.7), both generated by ETD of [L8S4 + 3H] (m/z 613.3), cf. Fig.

4.3b. Superscripted * or ‡ indicate fragments bound non-covalently to the intact PSS

chain (*) or a PSS chain that lost SO3H• (‡), respectively; the latter fragments have the

+ + compositions [bnS4 + H - SO3H•] and [ynS4 + H - SO3H•] . The sign # denotes

consecutive H2O or NH3 losses.

Our MS2 and MS3 findings clearly indicate that the non-covalent interactions between PLL and PSS are sufficiently strong to persist after a single CAD or ETD

34 activation event and are only disrupted by sequential ETD-CAD application. It is therefore theorized that PSS could potentially be an efficient and suitable “non-covalent label” for identifying basic surface-accessible sites of peptides and proteins by characterizing the sequence motif involved in their non-covalent interaction with PSS. This premise is evaluated with an MS2 and MS3 investigation of the non-covalent complex between PSS and substance P.

4.3.2. Substance P-PSS Complexes

Figure. 4. 7. (a) ESI-MS spectrum of the substance P + PSS mixture and (b) zoom-in

version of the m/z 960-1400 region. The doubly charged distribution of the substance

P-PSS complexes observed is marked on top of the corresponding peaks in red color. P

symbolizes substance P (undecapeptide with the sequence Arg-Pro-Lys-Pro-Gln-Gln- Phe-Phe-Gly-Leu-Met or R-P-K-P-Q-Q-F-F-G-L-M) and S the repeat unit of PSS; the

end groups of PSS have been omitted for brevity.

35 The ESI-MS spectrum acquired from solutions containing the undecapeptide substance P (R-P-K-P-Q-Q-F-F-G-L-M) and PSS shows a series of doubly protonated supramolecular complexes with the composition PSn (n = 3–6), where P designates substance P and Sn the non-covalently bound PSS chain, cf. Figure. 4. 7. The most abundant

2+ complex ion, viz. [PS4 + 2H] (m/z 1071.9) with four PSS repeat units, was selected for

MS2 and MS3 analysis.

2+ CAD of [PS4 + 2H] induces dissociations that are diagnostic for both the strength as well as binding sites of the non-covalent interaction between substance P and PSS

(Figure. 4. 8a). A partially contiguous series of bn* fragments (n = 5–10) with the intact

PSS chain is observed in the CAD spectrum, along with an abundant ion corresponding to substance P minus ammonia (m/z 665.8). The latter fragment provides compelling evidence that the non-covalent interaction between substance P and PSS is considerably weaker than that holding the PLL-PSS complex together. This observation is attributed to the presence of fewer potential binding sites on substance P whose sequence contains several amino acid residues with lower basicity compared to lysine.

In contrast to the CAD fragmentation pattern of substance P itself, which includes the formation of small an and bn fragments (n = 2–3, cf. Figure. 4. 9a), the smallest bn* fragment generated by CAD of the supramolecular bioconjugate is b5* (cf. Figure. 4. 8a).

This fragment contains the likely PSS binding sites Arg1, Lys3, and Gln5 which are located near the N-terminus, but not Gln6, which is another basic and, hence, potential binding site for an acidic polyelectrolyte. Our results strongly suggest that the former three amino acid residues are involved in the non-covalent interaction, whereas Gln6 is not, or is much more

weakly involved in such an interaction, presumably because its side chain extends in the opposite direction relative to those of Arg1, Lys3, and Gln5 in the substance P conformer sampled in the bioconjugation medium (see Experimental, section 4.2.1) [99]. Further validation of this assumption was sought by ETD and MS3 experiments.

2 2 2+ Figure. 4. 8. (a) MS (CAD) and (b) MS (ETD) spectra of [PS4 + 2H] (m/z 1071.9).

A superscripted asterisk (*) indicates fragments that contain the entire PSS chain, bound

+ non-covalently. Fragments cn* - 1 and [PS4 + H - SO3H2] are also present in the ETD

spectrum.

2+ ETD of [PS4 + 2H] yields a contiguous series of cn* fragments down to c5*, in striking similarity to CAD which yielded a continuous series of bn* fragments down to b5*

(Figure. 4. 8b vs. a). Under ETD conditions, however, the non-covalent interaction between

37 substance P and PSS is not broken, as no ions characteristic of detached substance P are observed. This difference between ETD and CAD is accounted for by the non-ergodic feature of the ETD process, which preserves the relatively labile non-covalent interaction between the undecapeptide and PSS [12,14].

Figure. 4. 9. (a) MS2 (CAD) spectrum of doubly protonated substance P, [P + 2H]2+

(m/z 674.9), and (b) MS2 (ETD) spectrum of triply protonated substance P, [P + 3H]3+

(m/z 450.3). These spectra, which reveal sequence information about Substance P, are

different from the corresponding spectra of substance P-PSS complex ions.

Compared to ETD of substance P, which causes fragmentation of essentially all N–

α C bonds to form almost complete cn and zn• fragment series (Figure. 4. 9), ETD of the PSS conjugate of this undecapeptide only produces the N-terminal cn* series comprising the intact PSS chain (Figure. 4. 8b), with the smallest fragment having five residues (c5*), in full agreement with the CAD results. This observation confirms that Gln6 may not be

38 involved in the non-covalent interaction between substance P and PSS, while the other three basic residues (Arg1, Lys3, and Gln5), which are closer to the N-terminus, are more likely to be the binding sites. Lastly, it is worth noting that a fragment arising by elimination of a SO3H• radical from PSS is also present in the ETD spectrum (Figure. 4. 8b); as discussed earlier, such a fragment corroborates the existence of non-covalent binding between a basic amino acid side chain and a –SO3H pendant on PSS (vide supra and

Scheme. 4. 1).

Sequential ETD-CAD was applied to c10* ions to verify the conclusions drawn from the single-stage ETD and CAD experiments. As expected, this fragment which represents the PSS conjugate of a truncated substance P (C-terminal Met is missing), and the original bioconjugate of intact substance P give rise to very similar CAD spectra, both showing a bn* fragment series down to b5*, cf. Figure. 4. 10 vs. Figure. 4. 8a. Furthermore, the non- covalent interaction between c10 and PSS is cleaved (Figure. 4. 10), as also observed for the bioconjugate between intact substance P and PSS (Figure. 4. 8a); this reconciles the

3 presence of unbound bn and internal c10yn fragments in the MS spectrum (Figure. 4. 10).

Overall, both the MS2 and MS3 data validate that Gln6 plays a minor role in the non-covalent interaction with PSS, as compared to Arg1, Lys3, and Gln5. This in turn indicates that Arg1, Lys3 and Gln5 must be the surface-accessible basic sites of substance

P that sense an approaching anionic polyelectrolyte like PSS, as also suggested by published NMR data [99].

3 Figure. 4. 10. MS (ETD-CAD) spectrum of c10* (m/z 2010.6), generated by ETD of

2+ [PS4 + 2H] (m/z 1071.4), cf. Fig. 4.8b. A superscripted asterisk (*) indicates fragments bound non-covalently to the PSS chain. The sign # denotes consecutive H2O or NH3 losses.

CHAPTER V

CONCLUSIONS

MS2 experiments on supramolecular PLL-PSS complexes, via either CAD or ETD, cause predominantly bond cleavages in the PLL backbone without disruption of the non- covalent interaction between PLL and PSS, which can only be broken by consecutive fragmentation via CAD of select ETD products. The ability to sequence peptide chains without affecting their non-covalent intermolecular binding to PSS reveals that PSS could act as an effectual “label” for determining the surface accessibility of basic residues on peptides or proteins by identification of the sequence motif containing the PSS label.

This premise was evaluated by elucidating the binding site of PSS in its supramolecular complex with substance P. Peptide fragments with the PSS chain were observed down to b5* (by CAD) or c5* (by ETD), indicating that amino acids up to residue

5 must be involved in the non-covalent binding of PSS and that Gln6 is not involved in the non-covalent interaction and, hence, is not an exposed site that can develop binding interactions with PSS. On the other hand, the participation of Arg1, Lys3, and Gln5 in the noncovalent binding of PSS is confirmed, since they are contained in all supramolecular fragments observed. Consequently, Arg1, Lys3, and Gln5 are the most likely surface- accessible sites of substance P. The non-covalent interaction between substance P and PSS is, however, weaker than that in the PLL-PSS complex, as it can be broken via single-stage

CAD.

41 The results reported in Chapter IV present a proof-of-concept for the effectiveness of PSS as a potential “non-covalent label” for the investigation of the surface accessibility of basic residues on biomolecules. This novel analytical concept can be extended to proteins to carry out surface mapping for more complicated structures.

42 REFERENCES

[1]. E.D. Hoffmann, V. Stroobant, Mass Spectrometry Principles and Applications,

2007.

[2]. T. Gruendling, S. Weidner, J. Falkenhagen, C. Barner-Kowollik, Mass

Spectrometry in Polymer Chemistry: A State-of-the-Art up-Date, Polym. Chem. 2010, 1,

599.

[3]. M. Karas, D. Bachmann, U. Bahr, Hillenkamp, F. Matrix-assisted ultraviolet laser desorption of non-volatile compounds, Int. J. Mass Spectrom. 1987, 78, 53-68.

[4]. M. Karas, F. Hillenkamp, Laser desorption ionization of proteins with molecular weights exceeding 10000 daltons, Anal. Chem. 1988, 60, 2299-2301.

[5]. K. Tanaka, H. Waki, Y. Ido, S. Akita, Y. Yoshida, T. Yohida, Protein and polymer analyses up to m/z 100 000 by laser ionization time-of-flight mass spectrometry, Rapid

Commun. Mass Spectrom. 1988, 2, 151–153.

[6]. M. Dole, L.L. Mack, R.L. Hines, R.C. Mobley, L.D. Ferguson, M.B. Alice,

Molecular beams of macroions, J. Chem. Phys. 1968, 49, 2240-2249.

[7]. J.S. Brodbelt, Ion Activation Methods for Peptides and Proteins, Anal. Chem. 2016,

88, 30–51.

[8]. T. Yalcin, W. Gabryelski, L. Li, Structural analysis of polymer end groups by electrospray ionization high-energy collision-induced dissociation tandem mass spectrometry, Anal. Chem. 2000, 72, 3847–3852.

43 [9]. K.M. Wollyung, C. Wesdemiotis, A. Nagy, J.P. Kennedy, Synthesis and mass spectrometry characterization of centrally and terminally amine-functionalized polyisobutylenes, J. Polym. Sci. Part A: Polym. Chem. 2005, 43, 946–958.

[10]. S.A. McLuckey, Principles of collisional activation in analytical mass spectrometry,

J. Am. Soc. Mass Spectrom. 1992, 3, 599-614.

[11]. L. Sleno, D.A. Volmer, Ion activation methods for tandem mass spectrometry, J.

Mass Spectrom. 2004, 39, 1091-1112.

[12]. R.A. Zubarev, N.L. Kelleher, F.W. McLafferty, Electron capture dissociation of multiply charged protein cations. A nonergodic process, J. Am. Chem. Soc. 1998, 120,

3265–3266.

[13]. R.A. Zubarev, D.M. Horn, E.K. Fridriksson, N.L. Kelleher, N.A. Kruger, M.A.

Lewis, B.K. Carpenter, F.W. McLafferty, Electron capture dissociation for structural characterization of multiply charged protein cations, Anal. Chem. 2000, 72, 563-573.

[14]. J.E.P. Syka, J.J. Coon, M.J. Schroeder, J. Shabanowitz, D.F. Hunt, Peptide and protein sequence analysis by electron transfer dissociation mass spectrometry, Proc. Nat.

Acad. Sci. 2004, 101, 9528–9533.

[15]. S.J. Pitteri, S.A. McLuckey, Recent developments in the ion/ion chemistry of highmass multiply charged ions, Mass Spectrom. Rev. 2005, 24, 931–958.

[16]. J.S. Brodbelt, Photodissociation Mass Spectrometry: New Tools for

Characterization of Biological Molecules, Chem. Soc. Rev. 2014, 43, 2757–2783.

[17]. J.P. Reilly, Ultraviolet Photofragmentation of Biomolecular Ions, Mass Spectrom

Rev. 2009, 28, 425–447.

44 [18]. T. Ly, R.R. Julian, Ultraviolet Photodissociation: Developments towards

Applications for Mass-Spectrometry-Based Proteomics, Angew. Chemie - Int. Ed. 2009,

39, 7130-7137.

[19]. F. Lermyte, D. Valkenborg, J.A. Loo, F. Sobott, Radical Solutions: Principles and

Application of Electron-Based Dissociation in Mass Spectrometry-Based Analysis of

Protein Structure, Mass Spectrom. Rev. 2018, 1, 1-22.

[20]. J.M. Wells, S.A. McLuckey, Collision-Induced Dissociation (CID) of Peptides and

Proteins, Methods Enzymol. 2005, 402, 143-185.

[21]. Y.H. Yang, K. Lee, K.S. Jang, Y.G. Kim, S.H. Park, C.S. Lee, B.G. Kim, Low

Mass Cutoff Evasion with Qz Value Optimization in Ion Trap, Anal. Biochem. 2009, 1,

133-135.

[22]. M.S. Kim, A. Pandey, Electron Transfer Dissociation Mass Spectrometry in

Proteomics, Proteomics. 2012, 7, 1625–1634.

[23]. L.M. Mikesh, B. Ueberheide, A. Chi, J.J. Coon, J.E.P. Syka, J. Shabanowitz, D.F.

Hunt, The Utility of ETD Mass Spectrometry in Proteomic Analysis, Biochimica et

Biophysica Acta - Proteins and Proteomics. 2006, 12, 1811-1822.

[24]. J. Wiesner, T. Premsler, A. Sickmann, Application of Electron Transfer

Dissociation (ETD) for the Analysis of Posttranslational Modifications, Proteomics. 2008,

21, 4466-4483.

[25]. S. Yin, J.A. Loo, Elucidating the site of protein-ATP binding by top-down mass spectrometry, J. Am. Soc. Mass Spectrom. 2010, 21, 899–907.

45 [26]. D.J. Clarke, E. Murray, T. Hupp, C.L. MacKay, P.R.R. Langridge-Smith, Mapping a noncovalent protein-peptide interface by top-down FTICR mass spectrometry using electron capture dissociation, J. Am. Soc. Mass Spectrom. 2011, 22, 1432–1440.

[27]. N.P. Barrera, C.V. Robinson, Advances in the mass spectrometry of membrane proteins: from individual proteins to intact complexes, Annu. Rev. Biochem. 2011, 80,

247–271.

[28]. J.P. O’Brien, W. Li, Y. Zhang, J.S. Brodbelt, Characterization of native protein complexes using ultraviolet photodissociation mass spectrometry, J. Am. Chem. Soc. 2014,

136, 12920–12928.

[29]. F. Chen, B. Gülbakan, S. Weidmann, S.R. Fagerer, A.J. Ibánez, ˜ R. Zenobi,

Applying mass spectrometry to study non-covalent biomolecule complexes, Mass

Spectrom. Rev. 2016, 35, 48–70.

[30]. H. Li, H.H. Nguyen, R.R. Ogorzalek Loo, I.D.G. Campuzano, J.A. Loo, An integrated native mass spectrometry and top-down proteomics method that connects sequence to structure and function of macromolecular complexes, Nat. Chem. 2018, 10,

139–148.

[31]. L. Sleno, D.A. Volmer, Ion activation methods for tandem mass spectrometry, J.

Mass Spectrom. 2004, 39, 1091–1112.[43][52]

[32]. Y. Xie, J. Zhang, S. Yin, J.A. Loo, Top-down ESI-ECD-FT-ICR mass spectrometry localizes noncovalent protein-ligand binding sites, J. Am. Chem. Soc. 2006,128, 14432–

14433.

46 [33]. K.O. Zhurov, L. Fornelli, M.D. Wodrich, Ü.A. Laskay, Y.O. Tsybin, Principles of electron capture and transfer dissociation mass spectrometry applied to peptide and protein structure analysis, Chem. Soc. Rev. 2013,42, 5014–5030.

[34]. J.J. Coon, Collision or electron? Protein sequence analysis on the 21st century, Anal.

Chem. 2009, 81, 3208–3215.

[35]. N.M. Riley, J.J. Coon, The role of electron transfer dissociation in modern proteomics, Anal. Chem. 2018, 90, 40–64.

[36]. K.F. Haselmann, T.J.D. Jørgensen, B.A. Budnik, F. Jensen, R.A. Zubarev, Electron capture dissociation of weakly bound polypeptide polycationic complexes, Rapid Commun.

Mass Spectrom. 2002, 16, 2260–2265.

[37]. S. Yin, J.A. Loo, Top-down mass spectrometry of supercharged native protein- ligand complexes, Int. J. Mass Spectrom. 2011, 300, 118–122.

[38]. D.L. Swaney, G.C. McAlister, M. Wirtala, J.C. Schwartz, J.E.P. Syka, J.J. Coon,

Supplemental activation method for high-efficiency electron-transfer dissociation of doubly protonated peptide precursors, Anal. Chem. 2007, 79, 477–485.

[39]. Z. Zhang, S.J. Browne, R.W. Vachet, Exploring salt bridge structures of gas-phase protein ions using multiple stages of electron transfer and collision induced dissociation, J.

Am. Soc. Mass Spectrom. 2014, 25, 604–613.

[40]. S. Gerislioglu, C. Wesdemiotis, Chain-end and backbone analysis of poly(N- isopropylacrylamide)s using sequential electron transfer dissociation and collisionally activated dissociation, Int. J. Mass Spectrom. 2017, 413, 61–68.

[41]. Y. Gao, S.A. McLuckey, Electron transfer followed by collision-induced dissociation (NET-CID) for generating sequence information from backbone modified oligonucleotide anions, Rapid Commun. Mass Spectrom. 2013, 27, 249–257.

[42]. B.C. Katzenmeyer, L.R. Cool, J.P. Williams, K. Craven, J.M. Brown, C.

Wesdemiotis, Electron transfer dissociation of sodium cationized polyesters: reaction time effects and combination with collisional activation and ion mobility separation, Int. J. Mass

Spectrom. 2015, 378, 303–311.

[43]. L. Vasicek, J.P. O’Brien, K.S. Browning, Z. Tao, H.-W. Liu, J.S. Brodbelt,

Mapping protein surface accessibility via an electron transfer dissociation selectively cleavable hydrazone probe, Mol. Cell. Proteomics 2012, 11, O111.015826.

[44]. D. Suckau, M. Mak, M. Przybylski, Protein surface topology-probing by selective chemical modification and mass spectrometric peptide mapping, Proc. Natl. Acad. Sci. U.

S. A. 1992, 89, 5630–5634.

[45]. M.O. Glocker, C. Borchers, W. Fiedler, D. Suckau, M. Przybylski, Molecular characterization of surface topology in protein tertiary structures by amino-acylation and mass spectrometric peptide mapping, Bioconjug. Chem. 1994, 5, 583–590.

[46]. F. Zappacosta, P. Ingallinella, A. Scaloni, A. Pessi, E. Bianchi, M. Sollazzo, A.

Tramontano, G. Marino, P. Pucci, Surface topology of Minibody by selective chemical modifications and mass spectrometry, Protein Sci. 1997, 6, 1901–1909.

[47]. B. Salih, R. Zenobi, MALDI mass spectrometry of dye-peptide and dye-protein complexes, Anal. Chem. 1998, 70, 1536–1543.

48 [48]. B.T. Turner Jr., T.M. Sabo, D. Wilding, M.C. Maurer, Mapping of factor XIII solvent accessibility as a function of activation state using chemical modification methods,

Biochemistry. 2004, 43, 9755–9765.

[49]. A. Leitner, W. Lindner, Functional probing of arginine residues in proteins using mass spectrometry and an arginine-specific covalent tagging concept, Anal. Chem. 2005,

77, 4481–4488.

[50]. A. Scholten, N.F.C. Visser, R.H.H. van den Heuvel, A.J.R. Heck, Analysis of protein-protein interaction surfaces using a combination of efficient lysine acetylation and nanoLC-MALDI-MS/MS applied to the E9:Im9 bacteriotoxin-immunity protein complex,

J. Am. Soc. Mass Spectrom. 2006, 17, 983–994.

[51]. V.L. Mendoza, R.W. Vachet, Protein surface mapping using diethylpyrocarbonate with mass spectrometric detection, Anal. Chem. 2008, 80, 2895–2904.

[52]. V.L. Mendoza, R.W. Vachet, Probing protein structure by amino acid-specific covalent labeling and mass spectrometry, Mass Spectrom. Rev. 2009, 28 785–815.

[53]. T. Liu, T.M. Marcinko, P.A. Kiefer, R.W. Vachet, Using covalent labeling and mass spectrometry to study protein binding sites of amyloid inhibiting molecules, Anal.

Chem. 2017, 89 11583–11591.

[54]. J.P. O’Brien, J.M. Pruet, J.S. Brodbelt, Chromogenic chemical probe for protein structural characterization via ultraviolet photodissociation mass spectrometry, Anal.

Chem. 2013, 85, 7391–7397.

[55]. J. Chen, D.L. Rempel, B.C. Gau, M.L. Gross, Fast photochemical oxidation of proteins and mass spectrometry follow submillisecond protein folding at the amino-acid level, J. Am. Chem. Soc. 2012, 45, 18724–18731.

49 [56]. Y. Pan, L. Konermann, Membrane protein structural insights from chemical labeling and mass spectrometry, Analyst. 2010, 135, 1191-1200.

[57]. L. Wang, M.R. Chance, Structural mass spectrometry of proteins using hydroxyl radical based protein footprinting., Anal. Chem. 2011, 83, 7234–7241.

[58]. X. Zheng, P.L. Wintrode, M.R. Chance, Complementary Structural Mass

Spectrometry Techniques Reveal Local Dynamics in Functionally Important Regions of a

Metastable Serpin, Structure. 2008, 16, 38-51.

[59]. K. Takamoto, M.R. Chance, Radiolytic protein footprinting with mass spectrometry to probe the structure of macromolecule complexes, Annu. Rev. Biophys.

Biomol. Struct. 2006, 35, 251−276.

[60]. S. Vahidi, B.B. Stocks, Y. Liaghati-Mobarhan, L. Konermann, Mapping pH- induced protein structural changes under equilibrium conditions by pulsed oxidative labeling and mass spectrometry, Anal. Chem. 2012, 84, 9124−9130.

[61]. Y. Pan, H. Piyadasa, J.D. O’Neil, L. Konermann, Conformational Dynamics of a

Membrane Transport Protein Probed by H/D Exchange and Covalent Labeling: The

Glycerol Facilitator, J. Mol. Biol. 2012, 416, 400−413.

[62]. P.A. Frantom, H.M. Zhang, M.R. Emmett, A.G. Marshall, J.S. Blanchard, Mapping of the Allosteric Network in the Regulation of α-Isopropylmalate Synthase from

Mycobacterium tuberculosis by the Feedback Inhibitor l-Leucine: Solution-Phase H/D

Exchange Monitored by FT-ICR Mass Spectrometry, Biochemistry. 2009, 48, 7457−7464.

[63]. V. Katta, B.T. Chait, S. Carr, Conformational changes in proteins probed by hydrogen‐exchange electrospray‐ionization mass spectrometry, Rapid Commun. Mass

Spectrom. 1991, 5, 214-217.

[64]. D. Rozbesky, P. Man, D. Kavan, J. Chmelik, J. Cerny, K. Bezouska, P. Novak,

Chemical Cross-Linking and H/D Exchange for Fast Refinement of Protein Crystal

Structure, Anal. Chem. 2012, 84, 867−870.

[65]. I.A. Kaltashov, C.E. Bobst, R.R. Abzalimov, H/D Exchange and Mass

Spectrometry in the Studies of Protein Conformation and Dynamics: Is There a Need for a

Top-Down Approach? Anal. Chem. 2009, 81, 7892−7899.

[66]. S. Valeja, M. Emmett, A. Marshall, Polar Aprotic Modifiers for Chromatographic

Separation and Back-Exchange Reduction for Protein Hydrogen/Deuterium Exchange

Monitored by Fourier Transform Ion Cyclotron Resonance Mass Spectrometry, J. Am. Soc.

Mass Spectrom. 2012, 23, 699−707.

[67]. Z. Zhang, D.L. Smith, Determination of amide hydrogen exchange by mass spectrometry: A new tool for protein structure elucidation, Protein Sci. 2008, 2, 522−531.

[68]. Z. Liu, R.R. Julian, Deciphering the Peptide Iodination Code: Influence on

Subsequent Gas-Phase Radical Generation with Photo dissociation ESI-MS, J. Am. Soc.

Mass Spectrom. 2009, 20, 965− 971.

[69]. J.K. Diedrich, R.R. Julian, Site-selective fragmentation of peptides and proteins at quinone-modified cysteine residues investigated by ESI-MS, Anal. Chem. 2010, 82, 4006–

4014.

[70]. Y. Tao, R.R. Julian, Examining protein surface structure in highly conserved sequence variants with mass spectrometry, Biochemistry. 2012, 51, 1796–1802.

[71]. W. Bleakney, A method of positive ray analysis and its application to the measurement of ionization potentials in mercury vapor, Phys. Rev. 1929, 34, 157-160.

51 [72]. A.O. Nier, A mass spectrometer for isotope and gas analysis, Rev. Sci. Instrum.,

1947, 18, 398-411.

[73]. A.G. Harrison, P. Lin, The chemical ionization mass spectra of fluorotoluenes, Can.

J. Chem. 1974, 53, 1314-1318.

[74]. A.G. Harrison, F.I. Onuska, C.W. Tsang, Chemical ionization mass spectrometry of specific polychlorinated biphenyl isomers, Anal. Chem. 1981, 53, 1183-1186.

[75]. D.I. Carroll, I. Dzidic, R.N. Stillwell, K.D. Haegele, E.C. Horning, Atmospheric pressure ionization mass spectrometry: Corona discharge ion source for use in liquid chromatograph-mass spectrometer-computer analytical system, Anal. Chem. 1975, 47,

2369-2373.

[76]. Z. Takats, J.M.Wiseman, B. Gologan, R.G. Cooks, Mass spectrometry sampling under ambient conditions with desorption electrospray ionization, Science. 2004, 306, 471-

473.

[77]. M.A. Grayson, John Bennett Fenn: a curious road to the prize, J. Am. Soc. Mass

Spectrom. 2011, 22, 1301-1308.

[78]. J.B. Fenn, M. Mann, C.K. Meng, S.F.Wong, C.M.Whitehouse, Electrospray ionization for mass spectrometry of large biomolecules, Science. 1989, 246, 64-71.

[79]. J.E. Campana, L.S. Sheng, S.L. Shew, B.E.Winger, Polymer analysis by photons, sprays, and mass spectrometry, Trends Anal. Chem. 1994, 13, 239-247.

[80]. S.D. Hanton, Mass spectrometry of polymers and polymer surfaces, Chem. Rev.

2001, 101, 527–570.

[81]. V.G. Zaikin, J.M. Halket, Derivatization in mass spectrometry—8. Soft ionization mass spectrometry of small molecules, Eur. J. Mass Spectrom. 2006, 12, 79-115.

52 [82]. B.E.Winger, K.J. Light-Wahl, R.R.O. Loo, H.R. Udseth, R.D. Smith, Observation and implications of high mass-to-charge ratio ions from electrospray ionization mass spectrometry, J. Am. Soc. Mass. Spectrom. 1993, 4, 536-545.

[83]. L. Konermann, E. Ahadi, A.D.Rodriguez, S. Vahidi, Unraveling the mechanism of electrospray ionization, Anal. Chem. 2013, 85, 2-9.

[84]. B.A. Thomson, J.V. Iribarne, Field induced ion evaporation from liquid surfaces at atmospheric pressure, J. Chem. Phys. 1979, 71, 4451-4463.

[85]. R.B. Cole, Some tenets pertaining to electrospray ionization mass spectrometry, J.

Mass Spectrom. 2000, 35, 763-772.

[86]. R. Knochenmuss, A quantitative model of ultraviolet matrix-assisted laser desorption/ionization including analyte ion generation, Anal. Chem. 2003, 75, 2199-2207.

[87]. W. Paul, H. Steinwedel, Ein neues Massenspektrometer ohne Magnetfel,

RZeitschrift für Naturforschung A. 1953, 8, 448-450.

[88]. R.G. Cooks, R.E. Keiser, Quadrupole ion trap mass spectrometry, Acc. Chem. Res.

1990, 23, 213-219.

[89]. G.L. Glish, Multiple stage mass spectrometry: The next generation tandem mass spectrometry experiment, Analyst. 1994, 119, 533-537.

[90]. T. Huang, S.A. McLuckey, Gas-phase chemistry of multiply charged bioions in analytical chemistry, Annu. Rev. Anal. Chem. 2010, 3, 365-385.

[91]. Compass Application Tutorial: ETD and PTR with the HCTultra, Version 1.3,

Bruker Daltonik GmbH, 2008, Bremen, Germany, pp. 9.

[92]. X.-J. Tang, R.K. Boyd, An investigation of fragmentation mechanisms of doubly protonated tryptic peptides, Rapid Commun. Mass Spectrom. 1992, 6, 651–657.

53 [93]. P. Wang, M.M. Kish, C. Wesdemiotis, Fragmentation mechanisms of peptide ions, in: R.S. Caprioli, M.L. Gross (Eds.), The Encyclopedia of Mass Spectrometry, Part A, vol.

2, Elsevier, Amsterdam. 2005, pp. 139–151, ISBN: 0080438008.

[94]. V. Scionti, C. Wesdemiotis, Tandem mass spectrometry analysis of polymer structures and architectures, in: C. Barner-Kowollik, in: T. Gruendling, J. Falkenhagen, S.

Weidner (Eds.), Mass Spectrometry in Polymer Chemistry, Wiley-VCH, Weinheim. 2012,

57–84.

[95]. C. Wesdemiotis, N. Solak, M.J. Polce, D.E. Dabney, K. Chaicharoen, B.C.

Katzenmeyer, Fragmentation pathways of polymer ions, Mass Spectrom. Rev. 2011, 30,

523–529.

[96]. J. Laskin, Surface-induced dissociation: a unique tool for studying energetics and kinetics of the gas-phase fragmentation of large ions, Eur. J. Mass Spectrom. 2015, 21,

377–389.

[97]. C. Cheng, M.L. Gross, Applications and mechanisms of charge-remote fragmentation, Mass Spectrom. Rev. 2000,19, 398–420.

[98]. M.J. Polce, M. Ocampo, R.P. Quirk, A.M. Leigh, C. Wesdemiotis, Tandem mass spectrometry characteristics of silver-cationized polystyrenes: internal energy, size, and chain end versus backbone substituent effects, Anal. Chem. 2008,80, 355–362.

[99]. G. Chassaing, O. Convert, S. Lavielle, Preferential conformation of substance P in solution, Eur. J. Biochem. 1986, 154, 77–85.

54 APPENDIX

55 56 57 58