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LYDIA R. COOL

SPECTROMETRY AND ION MOBILITY

A Dissertation

Presented to

The Graduate Faculty of The University of Akron

In Partial Fulfillment

of the Requirements for the Degree

Doctor of Philosophy

Lydia R. Cool

May, 2016 IDENTIFYING AND DISTINGUISHING ISOMERS USING MASS

SPECTROMETRY AND ION MOBILITY

Lydia R. Cool

Dissertation

Approved: Accepted:

______Advisor Department Chair Dr. Chrys Wesdemiotis Dr. Kim Calvo

______Committee Member Dean of the College Dr. Sailaja Paruchuri Dr. John Green

______Committee Member Dean of the Graduate School Dr. David Perry Dr. Chand Midha

______Committee Member Date Dr. Coleen Pugh

______Committee Member Dr. Claire Tessier

ABSTRACT

This dissertation focuses on the application of mass spectrometry (MS), tandem mass spectrometry (MS/MS), and ion mobility mass spectrometry (IM-MS) analysis of isomers. Chapter I gives an overview of the scope of the dissertation. Chapter II introduces mass spectrometry, including mass analyzers and ionization techniques.

Chapter III discusses the instrumentation and materials used in this dissertation.

Chapter IV discusses the analysis of five copolyesters. The first section of the chapter discusses two structural isomers synthesized using cyclohexane dicarboxylic acid

(CHDA) and either 1,5-pentanediol (1,5-PED) or neopentyl glycol (NPG), viz.

CHDA.NPG and CHDA.1,5-PED. Polyesters follow the 1,5-hydrogen rearrangement in

MS/MS experiments, but CHDA.NPG cannot dissociate via this mechanism. A distinct, charge-induced fragmentation mechanism is proposed to operate in this case based on

MS/MS fragmentation energetics and IM-MS results. The latter data serve a dual purpose as they can additionally be used to distinguish the two isomers. The second section of Chapter IV compares and contrasts the five isomers, which include oligomers made of adipic acid (AA) and ethylene glycol (1,2-EG). The hydrolysis behavior of the polyesters, which were synthesized by Mark D. Soucek et al. (University of Akron) are also discussed.

Chapter V reports the analysis of a fluorinated polymer. Sample preparation difficulties necessitated the use of a supercharging agent in order to analyze the sample

iii using ESI. Two different distributions were seen in the mass spectrum: linear and cyclic.

Despite multiple theoretical structures being possible, only one structure was confirmed present for either the linear or cyclic distributions. The cyclic structure was determined to be a complete circle (macrocycle), whereas the linear structure was found to be completely linear with no branching.

Chapter VI discusses multiple sets of monosaccharide-based isomers. Three different monosaccharides are included: α-D-mannopyranose, β-D-mannopyranose, and

β-D-glucopyranose. Seven different substituents were made onto the monosaccharides.

Tandem mass spectrometry cannot distinguish the isomers, but ion mobility mass spectrometry does. The IM-MS characteristics of the isomers followed a trend, independent of the substituent on the monosaccharides.

Chapter VII summarizes the conclusions of the dissertation. This is followed by the references. Finally are the appendices, which include copyright permissions.

iv DEDICATION

To my husband Elijah, my son Landon, and the rest of my wonderful family.

Your love and support has made each of these pages possible.

ACKNOWLEDGEMENTS

First and foremost, I would like to thank Dr. Wesdemiotis for many years of help and support. Starting in my undergraduate career, his assistance throughout my research and academic career has been invaluable. His kindness and guidance throughout the past six years is greatly appreciated.

I would like to thank my committee members: Dr. David Perry, Dr. Sailaja

Paruchuri, Dr. Coleen Pugh, and Dr. Claire Tessier. Thank you very much for your help and support throughout graduate studies, and specifically with this dissertation.

Thank you to the following group members: Vincenzo Scionti, Bryan

Katzenmeyer, Aleer Yol, Nadrah Alawani, Xiumin Liu, Ahlam Alawiat, Michelle

Kushnir, Sarah Robinson, Nick Alexander, Selim Gerislioglu, Sahar Sallam, Ivan Dolog,

Jailin Mao, and Savannah Snyder. Their assistance in the lab has been invaluable. I’d also like to thank my undergraduate student, Jordan Robideau, for his dedication to research and all his hard work.

I would like to thank Omnova Solutions Inc., particularly Dr. Matthew Espe, for the opportunity to gain professional experience by interning with them for two years.

The following collaborators have contributed greatly to the following work:

Cesar Lopez (Dr. Coleen Pugh, Department of Polymer Science), Matthew Quast (Dr.

Anja Mueller, Central Michigan University), and Mayela Ramirez-Huerta (Dr. Mark

Soucek, Department of Polymer Engineering). Additional collaborators have contributed

vi to my graduate studies, although our work is not included in this dissertation, and I greatly appreciate the experience these collaborations provided.

Last, but certainly not least, I would like to thank my family for their love and support. First of all, I would like to thank my husband, Elijah, for his unending dedication through these four years. I could not have finished this degree without your love and assistance. Second, to Landon, thank you for making every day an exciting adventure and for being the reason I worked so hard to finish. Third, I’d like to thank my parents, Crittenden and Carol Ohlemacher, for their support throughout my life and for encouraging me to follow my dreams and pursue higher education. Fourth, I would like to thank my mother-in-law Aimee Cool, for making me a member of her family from the very beginning and always taking an interest in my studies. Fifth, thank you to my sisters, Gwendolyn, Arielle, and Dominique, and brother-in-law, Nathan, for your support and your love. Lastly, to my in-laws, Chelsee, Joshua, and Granny, thank you for making me a part of your lives.

vii

TABLE OF CONTENTS

Page

LIST OF TABLES………………………………………………………………………..xi

LIST OF FIGURES……………………………………………………………………...xii

LIST OF SCHEMES…………………………………………………………………....xvi

LIST OF ABBREVIATIONS…………………………………………………………xviii

CHAPTER

I. INTRODUCTION…………………………………………………………………….1

II. INSTRUMENTAL METHODS AND BACKGROUND…………………………….5

2.1 Mass Spectrometry……………………………………………………………5

2.2 Ionization Techniques………………………………………………………...7

2.2.1 Electrospray ionization (ESI)……………………………………….7

2.2.2 Matrix-assisted laser desorption ionization (MALDI)……………...9

2.3 Mass Analyzers……………………………………………………………...11

2.3.1 Quadrupole Mass Analyzer (Q)…………………………………...11

2.3.2 Time-of-flight Mass Analyzer (ToF)……………………………...13

2.4 Detectors…………………………………………………………………….15

2.4.1 Daly detectors……………………………………………………..16

2.4.2 Microchannel plate detectors……………………………………...17

2.5 Tandem Mass Spectrometry (MS/MS)……………………………………...18

2.5.1 Definitions…………………………………………………………19

viii 2.5.2 Collisionally activated dissociation (CAD)……………………….20

2.5.3 Types of fragmentation……………………………………………20

2.6 Ion Mobility Mass Spectrometry (IM-MS)………………………………….21

2.6.1 Travelling wave ion mobility spectrometry (TWIMS)……...... 22

2.6.2 Collision cross section (CCS)……………………………………..24

III. MATERIALS AND INSTRUMENTATION………………………………………..26

3.1 Materials…………………………………………………………………….26

3.2 Instrumentation……………………………………………………………...27

3.2.1 Synapt HDMS……………………………………………………..27

3.2.2 Bruker Ultraflex III………………………………………………..28

IV. POLYESTERS……………………………………………………………………….30

4.1 Introduction………………………………………………………………….30

4.2 Experimental Method………………………………………………………..33

4.2.1 Synthesis of polyesters…………………………………………….33

4.2.2 MALDI preparation……………………………………………….34

4.2.3 ESI preparation……………………………………………………35

4.2.4 Synapt Q/ToF Parameters ………………………………………...35

4.2.5 Survival Yield Calculations……………………………………….35

4.3 Results……………………………………………………………………….37

4.3.1 Analysis of CHDA.NPG and CHDA.1,5-PED……………………37

4.3.2 Analysis of AA.1,2-EG, CHDA.1,2-EG, and AA.NPG…………..55

4.4 Conclusions………………………………………………………………….66

ix V. HYPERBRANCHED FLUORINATED POLYMERS……………………………...67

5.1 Introduction………………………………………………………………….67

5.2 Experimental Methods………………………………………………………68

5.2.1 Synthesis of Fluorinated Polymers………………………………..68

5.2.2 MALDI Preparation……………………………………………….68

5.2.3 ESI Preparation…………………………………………...... 69

5.2.4 Synapt Q/ToF Parameters ………………………………………...69

5.3 Results……………………………………………………………………….70

5.4 Conclusions………………………………………………………………….86

VI. SUGAR-BASED STRUCTURAL ISOMERS………………………………………88

6.1 Introduction………………………………………………………………….88

6.2 Experimental Methods……………………………………………………….90

6.2.1 Synthesis of the Fluorinated Polymer……………………………..90

6.2.2 ESI Preparation…………………………………………...... 90

6.2.3 Synapt Q/ToF Parameters…………………………………………90

6.3 Results………………………………………………………………………..91

6.4 Conclusions…………………………………………………………………108

VII. SUMMARY……………………………………………………………………….109

REFERENCES…………………………………………………………………………112

APPENDICES………………………………………………………………………….124

APPENDIX A. COPYRIGHT PERMISSIONS……………………………….125

APPENDIX B. DERIVATION OF E50 EQUATION………………………….128

x LIST OF TABLES

Table Page

4.1. Hydrolysis times of the various polyesters………………………………………34

4.2. The collision energies used for each oligomer of the five copolyesters…………35

4.3. Values of the coefficients and errors for the survival yield curves for CHDA.NPG and CHDA.1,5-PED……………………………………………………………...44

4.4. E50 values for CHDA.NPG and CHDA.1,5-PED oligomers…………………….46

4.5. Calibration Data………………………………………………………………….48

4.6. Experimental CCS for CHDA.NPG and CHDA.1,5-PED………………………49

4.7. Theoretical CCS for CHDA.NPG and CHDA.1,5-PED…………………………51

4.8. Mn values of the hydrolysis products of the five copolyesters.………………….58

4.9. Values of the coefficients and errors for the survival yield curves for AA.1,2-EG, CHDA.1,2-EG, and AA.NPG……………………………………………………63

4.10. E50 values for AA.1,2-EG, CHDA.1,2-EG, and AA.NPG oligomers……………64

5.1. Experimental CCS for cyclic and linear fluoropolymer oligomers………………...85

xi LIST OF FIGURES

Figure Page

2.1. A standard mass spectrometer. Modified from de Hoffmann.……………………6

2.2. Droplet formation and degradation in an ESI source. Reprinted with permission…………………………………………………………………………8

2.3. The principles of MALDI ionization…………………………………………….10

2.4. Quadrupole mass analyzer. Reproduced with permission………………………12

2.5. Stability of ions in the quadrupole. Reproduced with permission……………...13

2.6. Linear time-of-flight mass analyzer. Reproduced with permission……………..14

2.7. A reflection time-of-flight mass analyzer. Reproduced with permission……….15

2.8. A diagram of a Daly detector. Reproduced with permission……………………17

2.9. A microchannel plate. On top is a cross-section of the plate, and on bottom an example of the electron multiplication that occurs in a single channel. Reproduced with permission…………………………………………………….18

2.10. Traveling wave ion mobility spectrometry. Reproduced from article (ACS does not require permission for individual figures)…………………………………...22

2.11. Triwave portion of the Synapt HDMS mass spectrometer. Reproduced with permission………………………………………………………………………..23

3.1. Schematic diagram of the Synapt HDMS. Reproduced with permission……….27

4.1. MALDI spectra of CHDA.NPG after three hydrolysis times……………………38

4.2. MALDI spectra of CHDA.1,5-PED after three hydrolysis times………………..39

4.3. MS/MS spectrum of the sodiated 3-mer (1085 m/z) of CHDA.1,5-PED. Monoisotopic m/z values are given on select peaks. The m/z values of the fragments formed by consecutive dissociations are underlined…………………40

xii 4.4. MS/MS spectra of the sodiated 3-mer of CHDA.NPG (1085 m/z). Monoisotopic m/z values are given of select peaks. The m/z values of the fragments formed by consecutive dissociations are underlined………………………………………...43

4.5. Survival yield curves for sodiated CHDA.NPG oligomers……………………...45

4.6. Survival yield curves for sodiated CHDA.1,5-PED oligomers………………….45

4.7. IM-MS plot (m/z vs. drift time) of CHDA.1,5-PED. The [M+Na]+ ions of intact oligomers are encased. The number of repeat units is also given…………….…47

4.8. IM-MS plot (m/z vs. drift time) of CHDA.NPG. The [M+Na]+ ions of intact oligomers are encased. The number of repeat units is also given……………….47

4.9. Calibration curve for ubiquitin…………………………………………………...48

4.10. Calculated CCS vs. relative energy for 150 energy-minimized structures of the CHDA.1,5-PED 2-mer. The three different sets of 50 are designated by the three different colors…………………………………………………………………...50

4.11. Representative energy-minimized structures for the sodiated 2-mers from (a) CHDA.NPG and (b) CHDA.1,5-PED……………………………………………52

4.12. MALDI mass spectra of AA.1,2-EG after three hydrolysis times……………….55

4.13. MALDI mass spectra of CHDA.1,2-EG after three hydrolysis times…………...56

4.14. MALDI mass spectra of AA.NPG after two hydrolysis times…………………..57

4.15. The MS/MS spectrum of the sodiated CHDA.1,2-EG 4-mer acquired at a CE of 55 eV……………………………………………………………………………..59

4.16. The MS/MS spectrum of the sodiated AA.1,2-EG 4mer acquired at a CE of 45 eV………………………………………………………………………………...60

4.17. The MS/MS spectrum of the sodiated AA.NPG 3-mer acquired at a CE of 65 eV………………………………………………………………………………...61

4.18. Survival yield curves for sodiated AA.1,2-EG oligomers……………………….62

4.19. Survival yield curves for sodiated CHDA.1,2-EG oligomers……………………62

4.20. Survival yield curves for sodiated AA.NPG oligomers…………………………...63

5.1. MALDI mass spectrum of the fluorinated polymer. All peaks correspond to [M+Ag]+ ions…………………………………………………………………….71

xiii 5.2. Zoomed-in view of MALDI spectrum showing the two distributions. All peaks correspond to [M+Ag]+ ions……………………………………………………..72

5.3. ESI spectrum of the fluorinated polymer. Ln designates oligomers still containing the OH focal point (structure shown in Scheme 5.1). Cn designates cyclic oligomers (macrocycles or tadpoles, see text)…………………………………...73

5.4. MS/MS spectrum of the [M+Ag]+ ion from the 2-mer of the linear fluorinated polymer, acquired at a CE of 40 eV……………………………………………...74

5.5. IM-MS spectra for (a) cyclic 3-mer and (b) linear 3-mer………………………..77

5.6. MS/MS spectrum of the silverated 3-mer from the cyclic distribution of the fluorinated polymer, acquired at a collision energy of 55 eV……………………79

5.7. MS/MS spectrum of the silverated 3-mer from the linear distribution of the fluorinated polymer. (CE = 55 eV)………………………………………………80

5.8. MS/MS spectrum of the silverated 4-mer from the cyclic distribution of the fluorinated polymer (CE = 60 eV)……………………………………………….81

5.9. Calibration curve for ubiquitin…………………………………………………...86

6.1. Mass spectrum of R1-substitued α-mannose. Calculated mass = 413.1054 Da. Mass accuracy = 123 ppm……………………………………………………….91

6.2. Mass spectrum of R2-substituted α-mannose. The peak at m/z 449 arises from an impurity; the measured m/z corresponds to a sodiated mannose disaccharide with two acetyl groups (C16H26O13). Calculated mass = 447.0665 Da. Mass accuracy = 127 ppm……………………………………………………….92

6.3. Mass spectrum of R3-substituted α-mannose. Calculated mass = 427.1211 Da. Mass accuracy = 217 ppm……………………………………………………….93

6.4. Mass spectrum of R4-substituted α-mannose. Calculated mass = 425.1054 Da. Mass accuracy = 226 ppm……………………………………………………….93

6.5. Mass spectrum of α-mannose of the R5 substituted sugar. Calculated mass = 475.1211 Da. Mass accuracy = 153 ppm………………………………………..94

6.6. Mass spectrum of α-mannose of the R6 substituted sugar. Calculated mass = 491.1524 Da. Mass accuracy = 219 ppm………………………………………..94

6.7. Mass spectrum of α-mannose of the R7 substituted sugar. Calculated mass = 885.2999 Da. Mass accuracy = 181 ppm………………………………………..95

xiv 6.8. MS/MS spectra of the R3-substituted monosaccharides. CE = 22 eV………….96

6.9. IM-MS spectra of the R1-substituted monosaccharides: (a) α-D-mannopyranose, (b) β-D-mannopyranose, and (c) β-D-glucopyranose……………………………98

6.10. IM-MS spectra of the R2-substituted monosaccharides: (a) α-D-mannopyranose, (b) β-D-mannopyranose, and (c) β-D-glucopyranose……………………………99

6.11. IM-MS spectra of the R3-substituted monosaccharides: (a) α-D-mannopyranose, (b) β-D-mannopyranose, and (c) β-D-glucopyranose…………………………..100

6.12. IM-MS spectra of the R4-substituted monosaccharides: (a) α-D-mannopyranose and (b) β-D-mannopyranose……………………………………………………101

6.13. IM-MS spectra of the R5-substituted monosaccharides: (a) α-D-mannopyranose, (b) β-D-mannopyranose, and (c) β-D-glucopyranose…………………………..102

6.14. IM-MS spectra of the R6-substituted monosaccharides: (a) α-D-mannopyranose, (b) β-D-mannopyranose, and (c) β-D-glucopyranose………………………...... 103

6.15. IM-MS spectra of the R7-substituted monosaccharides: (a) α-D-mannopyranose, (b) β-D-mannopyranose, and (c) β-D-glucopyranose…………………………..104

6.16. The IM-MS spectra of the R1-substituted monosaccharides: (a) α-D- mannopyranose, (b) β-D-mannopyranose, (c) β-D-glucopyranose, and (d) the mixture of the three……………………………………………………………..105

6.17. The IM-MS spectra of the R2-substituted monosaccharides: (a) α-D- mannopyranose, (b) β-D-mannopyranose, (c) β-D-glucopyranose, and (d) the mixture of the three……………………………………………………………..106

6.18. The IM-MS spectra of the R7-substituted monosaccharides: (a) α-D- mannopyranose, (b) β-D-mannopyranose, (c) β-D-glucopyranose, and (d) the mixture of the three……………………………………………………………..107

xv LIST OF SCHEMES

Scheme Page

2.1. Two different fragmentation mechanisms……………………………………….20

4.1. The condensation/hydrolysis reaction. X is the R group of the diol. Y is the R group of the diacid……………………………………………………………….30

4.2. The five copolyesters to be discussed in this Section……………………………31

4.3. The 1,5-hydrogen rearrangement, as it applies to copolyesters…………………32

4.4. 1,5-Hydrogen rearrangement in collisionally activated CHDA.1,5-PED……….40

4.5. Phenyl isocyanate loss from the copolyester end groups………………………..41

4.6. Major and minor fragments in the MS/MS spectrum of sodiated CHDA.1,5-PED trimer…………………………………………………………………………….42

4.7. CHDA.1,5-PED fragment end group structures………………………………....43

4.8. Proposed fragmentation mechanism for the COO-CH2 bond cleavage in sodiated CHDA.NPG……………………………………………………………………...53

4.9. CHDA.NPG fragment end group structures……………………………………..54

5.1. Polymerization of Fluorinated Polymer………………………………………….68

5.2. Plausible structures of the MS/MS fragments from the linear 2-mer. Isomeric structures are possible……………………………………………………………75

5.3. Two possible structures for the cyclic 3-mer…………………………………….78

5.4. Structurally diagnostic MS/MS fragments expected from the linear and branched architecture of the 3-mer. Note that the linear structure has positional isomers, formed by chain propagation at the pentafluorophenyl rings attached at C3 of the central phenyl group. All bond clevages shown (except the 181 Da loss) are accompanied by H-rearrangement to the moiety being lost……………………..83

xvi 5.5. Structurally diagnostic MS/MS fragments from the 4-mer with linear architecture……………………………………………………………………….84

6.1. Monosaccharides examined for incorporation into glycopolymers……………...89

6.2. Monosaccharide derivatives studied……………………………………………..89

6.3. Fragmentation pathways accounting for the MS/MS fragments from the R3- substituted sugar monosaccharide………………………………………………..97

xvii LIST OF ABBREVIATIONS

AgTFA silver trifluoroacetate

ACN acetonitrile

CAD collision activated dissociation

CDEM continuous-dynode electron multiplier

CID collision induced dissociation

CCS collisional cross section

DCTB trans-2-[3-(4-tert-butylphenyl)-2-methyl-2-propenylidene]malononitrile

DMSO dimethyl sulfoxide

DT drift time

DT’ corrected drift time

E50 ECM value when SY = 0.5

Ecm energy of center of mass

Elab laboratory frame energy

EI electron ionization

ESI electrospray ionization

GC gas chromatography

HPLC high performance liquid chromatography

IT ion trap

LC liquid chromatography

xviii Mg mass of the collision gas

Mi mass of the precursor ion

Mp mass of molecule without salt

MALDI matrix-assisted laser desorption ionization

MeOH methanol

MS mass spectrometry

MS/MS tandem mass spectrometry

MSn multidimensional tandem mass spectrometry m/z mass to charge ratio

NaTFA sodium trifluoroacetate

Q quadrupole

QIT quadrupole ion trap

Q/ToF quadrupole/time-of-flight

SY survival yield

THF tetrahydrofuran

ToF time-of-flight

TWIMS travelling wave ion mobility spectrometry z charge of the ion

Ω CCS

Ω’ corrected CCS

μ reduced mass

xix

CHAPTER I

INTRODUCTION

Mass spectrometry (MS) is a powerful analytical technique used to analyze a wide range of compounds, from polymers to biological samples to inorganics. Mass spectrometry has advantages over other analytical techniques in that it requires very little sample, can analyze low concentration contaminates, and is capable of distinguishing mixtures. MS measures mass-to-charge ratios (m/z) of the compound. Mass spectrometers follow the same five components: an inlet system, ionization chamber, mass analyzer, ion detector, and data system. Some common ionization techniques are matrix-assisted laser desorption ionization (MALDI) and electrospray ionization (ESI).

In MALDI, a laser is used to irradiate a mixture of sample and matrix. This causes the sample to enter the gas phase and be ionized. Vitally important to this method is the matrix, which is a small molecule that absorbs at the wavelength of the laser. This matrix protects the sample from being destroyed by the laser, while assisting in ionization. This technique is particularly powerful as it can be used to ionize intact high molecular weight samples.

In ESI, the sample is dissolved in a volatile solvent and a syringe is used to spray the solution into the ionization chamber. A heated desolvation gas is used to remove the solvent. Individual droplets of sample and ions experience charge repulsion until they reach the Rayleigh limit and smaller droplets are formed. This continues until individual

1 analytes remain with one or more charges on the surface. There are many advantages to

ESI, including the possibility of multiple charges. This means that analytes of higher molecular weight than the mass limit of the instrument can be analyzed, including high molecular weight proteins and biomolecules.

Whereas mass spectrometry has many advantages, there are some disadvantages.

There are limitations, even with MALDI, of how high of a molecular weight can be analyzed. ESI in is a liquid technique, which requires the sample to follow certain solubility guidelines. However, one of the largest disadvantages, and the focus of this dissertation, is mass spectrometry’s inability to distinguish isomers. Since isomers have the same molecular weight, they cannot be separated by standard mass spectrometry alone. This requires the introduction of multidimensional mass spectrometry techniques or “hyphen” techniques. These included tandem mass spectrometry (MS/MS), liquid chromatography (LC), gas chromatography (GC), and ion mobility mass spectrometry

(IM-MS).

Tandem mass spectrometry (MS/MS) can be useful in determining structural information about a molecule. The molecular ion, or parent ion, is fragmented using collisionally activated dissociation (CAD) or collision-induced dissociation (CID). The fragments are then analyzed using a second mass analyzer. These fragments provide clues to the structure of the molecule. MS/MS has been particularly advantageous for determining end groups and architectures of polymers. In addition, survival yield curves can be calculated using MS/MS, which provides a characteristic value called the E50 value, which can be used to compare and identify analytes.

2 Ion mobility mass spectrometry (IM-MS) separates ions based on their shape and size, rather than by their mass. This makes this technique particularly advantageous for analyzing isomers. In this technique, the ions enter the ion mobility cell and are pushed along by an electric field while also colliding with a buffer gas. The smaller the ion, the more easily it will flow through the gas. In addition, a molecule with the same size but different charges will interact differently with the electric field. IM-MS data can be used to calculate collision cross sections (CCS). This value is characteristic to the sample, and when compared to theoretical structures, can be used to determine architectures.

This dissertation focuses on the application of mass spectrometry to the analysis of isomers. In Chapter II, the basics of mass spectrometry will be discussed, including ionization techniques, mass analyzers, tandem mass spectrometry, and ion mobility.

Chapter III will discuss the materials and methods used in this dissertation. Chapters IV

– VI deal with the analysis of three different isomer sets. Chapter VII summarizes the conclusions from each of the research chapters.

Chapter IV discusses the analysis of five copolyesters, two of which are structural isomers. Mayela Cristina Ramirez-Huerta, from Dr. Soucek’s research group, synthesized the samples. The tandem mass spectra for the two isomers are identical, which led to two research questions: first, how can the spectra be identical when the isomers cannot follow the same fragmentation mechanism, and second, how can mass spectrometry be used to distinguish the two isomers if MS/MS will not work? In order to distinguish the isomers, collisional cross sections (CCS) and survival yield (SY) curves were calculated. This data was used to propose a separate fragmentation mechanism for

3 one the isomers. In addition, comparisons were made between the five copolyesters and trends were studied.

Chapter V discusses the analysis of a hyperbranched fluorinated polymer.

Matthew Quast, from Central Michigan University, synthesized the samples. The hydrophobicity of the sample complicated the analysis and required special sample preparation. Two distributions are present in the sample: linear and cyclic. The structure will be determined for both of these distributions using ion mobility mass spectrometry.

Chapter VI discusses the analysis of sugar-based structural isomers. These compounds are of particular interest, as they can be used for biomimetic polymers.

Seven different substitutions were made onto three different sugars: α-D-mannopyranose,

β-D-mannopyranose, and β-D-glucopyranose. Cesar Lopez, from Dr. Pugh’s research group, synthesized these samples. The different substitutions were confirmed using mass spectrometry. The isomers were distinguished using ion mobility and a mixture of the isomers was studied.

CHAPTER II

INSTRUMENTAL METHODS AND BACKGROUND

2.1 Mass Spectrometry

In 1886, J.J. Thomson discovered that when ions were formed using a gas discharge apparatus and electric and magnetic fields were applied, the ions formed parabolic trajectories.1 It was found that these parabolas represented different mass-to-

+ + 2 charge ratios, and this instrument could distinguish between H and H2 . When analyzing a neon sample, the standard parabola for Ne corresponding to the atomic weight of 20 was seen, as well as an additional parabola that was determined to be at an

2 atomic weight of 22. This mass at 22 was initially thought to be NeH2, but eventually proved to be an isotope of Ne, 22Ne. Based on the principle of isotope separation, the first mass spectrometer (at the time called a mass spectrograph) was built in 1907 by

Thomson in Cavendish Laboratory at Cambridge University.2 The ability of mass spectrometry to separate isotopes was proven to be a powerful feature for future applications of the technique.

Mass spectrometers follow the same general layout (Figure 2.1). The sample is introduced into the mass spectrometer by an inlet. This inlet could be from a gas chromatograph (GC), liquid chromatograph (LC), or direct injection port, the primary method for this dissertation. Direct injection occurs when the sample is directly inserted to the ionization source, either by a syringe pump or by inserting the solid sample on a

5 plate. After injecting the sample, it is ionized in the ionization source using one of many techniques, which will be further discussed in Section 2.2. The sample then enters into the first mass analyzer. Mass analyzers will be discussed in Section 2.3. If the sample is being analyzed using standard mass spectrometry, or one-dimensional mass spectrometry, the first mass analyzer and fragmentation chamber are not active and the sample passes directly to the second mass analyzer.

Figure 2.1. A standard mass spectrometer.

Modified from de Hoffmann.3

If the sample is being analyzed using tandem mass spectrometry (MS/MS), the parent (or precursor) ion is selected by the first mass analyzer, and then sent to the fragment chamber. The fragments formed in this chamber are sent to the second mass analyzer.

Tandem mass spectrometry will be discussed in greater length in Section 2.5. The ions are then sent to the detector, which will be discussed in Section 2.4. In some cases, additional chambers exist in the mass spectrometer, such as an ion mobility chamber, which will be discussed in Section 2.6.

6 2.2 Ionization Techniques

One of the first ionization techniques was electron ionization (EI).3 EI was first introduced in 1929 by Bleakney4 and further revised in 1947 by Nier.5 In EI, a beam of electrons collides with the sample, producing ions, which are further analyzed in the mass spectrometer. While still a popular ionization method today, EI is considered a hard ionization method, in that it induces fragmentation of the parent ion. Softer techniques, such as electrospray ionization (ESI) and matrix-assisted laser desorption ionization

(MALDI), were invented at a later date and will be discussed below.

2.2.1 Electrospray Ionization (ESI)

Although it had existed previously, ESI’s prominence as an ionization source truly emerged when Mann et al. discovered proteins can be multiply charged using the

ESI source.6 Due to this discovery, analyzers with a detection limit as low as 2000 Da could still have practical applications.3 For example, a protein of molecular weight of 50 kDa could be analyzed by a mass spectrometer with a mass limit of 4000 Da if the protein can be multiply charged to a minimum of 13 charges. At first, ESI sources were primarily used for proteins, but their potential applications expanded to include polymers and small molecules.7–11

In order to create an ESI spectrum, a strong electric field is applied to the liquid passing through a capillary under atmospheric pressure, typically at a low flow, such as 1

– 10 uL/min.3,12 An electric potential difference is applied between the capillary and the lens at the entrance of the vacuum system, which causes the solution to spray from the capillary towards the lens (Figure 2.2). Specifically, this potential difference leads to a

7 charge accumulation at the liquid surface at the end of the capillary. This surface charge leads to the expulsion of highly charged droplets, forming a Taylor cone.3 The presence of a Taylor cone in ESI was first noted by Gomez and Tang.13 A heated neutral gas (such as N2) is applied perpendicular to the spray, which causes the solvent to evaporate from the droplets. This causes a charge build up on the surface of the droplet, until the droplet reaches its Rayleigh limit, at which point the droplet is further reduced in size. This process continues until there is a single molecule with charges on the surface (Figure

2.2).3,12 These ions then enter the analyzer.

Figure 2.2. Droplet formation and degradation in an ESI source.

Reprinted with permission.14

ESI has several advantages. As a soft ionization technique, intact parent ions are analyzed and detected.12 In addition, because ESI has the ability to form multiply charged ions, this technique can analyze larger molecules than would normally be visible in the mass analyzer.3 There are potential problems with the spraying of the solvent

(depending on the onset voltage necessary to see ionization with the solvent) that can

8 limit the samples that can be analyzed. ESI is also very sensitive to concentration, as well as salts and buffers and other contaminants.

2.2.2 Matrix-Assisted Laser Desorption Ionization (MALDI)

Karas and Hillenkamp first introduced MALDI instruments in the late 1980s.3

MALDI is a soft ionization technique that generates intact gas-phase ions. In order to analyze the sample, it is combined with a matrix (which will be explained in greater length below). A salt is added, if necessary, and two possible preparation methods are utilized. The first, the sandwich method, uses a solution containing the matrix and salt and a separate solution containing the sample. The matrix/salt solution is added to the

MALDI plate and the solvent allowed to evaporate, then the sample solution is applied to the plate and the solvent allowed to evaporate, and then a final layer of the matrix/salt solution is applied. The second method, the dry-drop method, combines the matrix, sample, and salt into one solution, which is applied to the MALDI plate, and the solution allowed to dry. When the solvent evaporates, a ‘solid solution’ of the sample, matrix, and salt is left on the plate, as seen in Figure 2.3. The plate is then inserted in the

MALDI source.

9 Figure 2.3. The principles of MALDI ionization.

The ‘solid solution’ is bombarded with a laser, which causes the matrix to desorb off the plate and into the gas phase.3 The matrix is then removed from the sample through a desolvation process and the charged sample continues into the mass analyzer.

The properties of the matrix make MALDI possible.3 Matrices are small, aromatic compounds that absorb at a particular wavelength. This wavelength is in the spectrum of the laser, which excites the matrix. This causes a local increase in heat that desorbs the matrix off the plate, which takes the intact sample along in the matrix plume.

(The mechanism of this process is not exactly understood.) The matrix is in large excess of the sample, which completely isolates the sample molecules from each other. The matrix, when desorbed with the laser, can protonate or deprotonate the sample to ionize it.

10 There are certain advantages to MALDI. It is not as sensitive to contaminants, such as salts or buffers.3 The pulsed nature of the MALDI source makes it particularly useful for ToF analysis. MALDI can measure higher molecular weights than ESI, with some MALDI mass spectrometers capable of measuring proteins with a molecular weight of 300 kDa.3

2.3 Mass Analyzers

After forming gas phase ions in the ionization source, the ions need to be separated by their mass-to-charge ratios (m/z). This is accomplished in a mass analyzer.

Mass analyzers are held under vacuum to prevent interactions between the gas phase ions and any gaseous molecules in the environment. A wide variety of mass analyzers exist, including: magnetic sector, quadrupole (Q), ion trap (IT), and time-of-flight (ToF) mass analyzers.3 Each of these analyzers has different strengths and weaknesses. For example, magnetic sector instruments can provide high mass accuracy and excellent resolution;15 however, the energy required to supply the magnetic field and keep it constant can be prohibitively expensive. In addition, the magnetic portion of the instrument can be quite large.

Two different mass analyzers were used for the work in this dissertation and will be discussed in the following sections.

2.3.1 Quadrupole Mass Analyzer (Q)

First suggested in 1953, quadrupoles are perfectly parallel rods that are typically circular.3 The rods are arranged in a square configuration, with the two rods in opposite

11 corners being similarly charged (i.e. positively charged) and the other two corners being oppositely charged (i.e. negatively charged) (Figure 2.4). The rods change charge as a function of time.

Figure 2.4. Quadrupole mass analyzer.

Reproduced with permission.16

A positively charged ion will be attracted towards the negatively charged rod. If the ion reaches the rod before the rod changes polarity, it will touch the rod and become neutral.

Since mass spectrometry can only analyze ions, these neutral molecules will not be analyzed. (This is how quadrupoles can selectively analyze a particular mass range, i.e. 0

– 8kDa.) However, if the polarities change before the ion reaches the rod, it will move towards the negative rod closest to its location. The zigzag motion will continue until the ion exits the quadrupole (stable path). Both a stable and unstable ion path can be seen in

Figure 2.5.

12 Figure 2.5. Stability of ions in the quadrupole.

Reproduced with permission.3

Quadrupoles are commonly used as the first mass analyzer when multiple mass analyzers are combined in the same mass spectrometer.

2.3.2 Time-of-flight Mass Analyzer (ToF)

Time-of-flight analyzers were first discussed by Stephens in 1946.3,17 ToF is particularly suited for use with laser desorption ionization methods, such as MALDI due to the pulsed nature of that ionization source. ToF analyzers include both linear and reflectron time-of-flight analyzers. Each of these will be discussed below.

13 Linear time-of-flight analyzers were first designed in 1955 by Wiley and

McLauren.18 An image of a linear ToF coupled with a MALDI source can be seen in

Figure 2.6.

Figure 2.6. Linear time-of-flight mass analyzer.

Reproduced with permission.3

The ions are first accelerated by electric fields in what the figure calls the acceleration region. These electric fields apply a certain amount of kinetic energy (KE) to each ion.

The ions then leave the field and enter the field-free flight tube of length L. Because the ions have a constant kinetic energy, the ions will have a characteristic velocity as a function of their mass (Equation 2.1).

KE = ½mv2 Equation 2.1

This separates the ions in the tube by mass. The ions continue down the field-free region until they reach the detector.3

Reflectron time-of-flight analyzers were invented to improve the mass resolution of the linear ToF.19 With linear ToF instruments, slight differences in the kinetic energy

14 applied to individual ions would cause peak broadening. This problem can be resolved by adding an electric field to the end of the field-free region (Figure 2.7).3

Figure 2.7. A reflection time-of-flight mass analyzer.

Reproduced with permission.3

The ions enter the electric field and are reflected back down the field-free region, hence the name reflectron ToF. The ions with higher velocities (and therefore higher kinetic energies) will enter deeper into the reflectron region, whereas those with lower velocities will not be able to penetrate as far. This variation corrects for the energy errors, and allows ions of the same mass to reach the detector at the same time, which improves the overall resolution.

2.4 Detectors

The function of the detector is to detect and transform the presence of the ion into data usable by the output computer.3 Detectors create an electrical current that is proportional to the ion abundance.3 While new research into ionization methods and mass analyzers occurs regularly, little research has been done to update the detectors used

15 in mass spectrometry.20 Two detectors will be covered in this section: Daly and microchannel plate detectors.

2.4.1 Daly Detectors

The Daly detector is a combination of ion and photon detection devices, which convert ions to electrons and then photons (Figure 2.8).3,20 The Daly detector is made up of two different parts: the phosphorescent screen and the photomultiplier. Detection begins when the ions from the mass analyzer strike a dynode. A dynode converts an input ion, either positive or negative, into electrons.3 The electrons are accelerated toward the phosphorescent screen, which converts the electrons into photons, which are then sent to the photomultiplier. This produces an electric current, which is amplified and sent to the computer.3

Figure 2.8. A diagram of a Daly detector.

Reproduced with permission.3

There are multiple advantages to using the multistage electro-optical ion detector.3 It has a longer lifetime than standard detectors, because it is sealed in glass under vacuum. This detector has a fast response time and high sensitivity, which can amplify signals 104 to

105 times.3

2.5.1 Microchannel plate detectors

A microchannel plate detector is a version of an electron multiplier, the most widely used ion detector (Figure 2.9).3 Specifically, it is a version of a continuous- dynode electron multiplier (CDEM).20 In this detector, a dynode is used to convert the

17 ion into electrons, once it strikes the side of the channel. These electrons then continue down the channel, striking the side again, creating more electrons.

Figure 2.9. A microchannel plate. On top is a cross-section of the plate, and on bottom

an example of the electron multiplication that occurs in a single channel.

Reproduced with permission.3

This cascading effect continues down the length of the channel, increasing the intensity of the signal by up to 108 fold.3 Microchannel plates are particularly effective with ToF instruments, as the plates are extremely sensitive to the arrival time of the ion and have narrow pulse widths.20 However, these plates can be fragile and sensitive to their environment.20

2.5 Tandem Mass Spectrometry (MS/MS)

Tandem mass spectrometry (MS/MS) employs two stages of mass analysis with fragmentation of the ions occurring between the two stages.3 There are two types of fragmentation: in space and in time.3 In space fragmentation occurs when the ions pass through one mass analyzer, enter some sort of collision cell and are fragmented, and then

18 the fragmented ions enter a second mass analyzer where they are analyzed. Typical in space instrumentation includes a quadrupole/time-of-flight (Q/ToF) and ToF/ToF instruments. In time fragmentation occurs specifically in ion trap mass analyzers, where the parent ion is selected and stored in the ion trap and then fragmentation occurs during a certain time period. The ions are then ejected and analyzed. Ion traps permit multidimensional tandem mass spectrometry (MSn) experiments by repeating the steps of ejection and fragmentation.

2.5.1 Definitions

The following definitions are necessary in order to discuss MS/MS.21,22 The precursor or parent ion is the ion that undergoes fragmentation. The product or daughter ion(s) is/are the ion(s) that result(s) from the fragmentation. Collisionally activated dissociation (CAD) occurs when a neutral gas is used to induce the fragmentation. This is also known in the literature as collision induced dissociation (CID). Neutral losses occur when the fragmentation results in a neutral fragment, which cannot be analyzed by mass spectrometry. Charge induced fragmentation occurs if fragmentation is induced by the charge on the parent ion.23 Charge remote fragmentation occurs when the fragmentation location is independent of the location of the charge.

+ + Mp → Mf + Mn

+ The equation above shows the relationship between the precursor ion (Mp ), product ion

+ (Mf ), and neutral losses (Mn).

19 2.5.2 Collisionally activated dissociation (CAD)

CAD occurs when the precursor ion collides with a neutral gas, such as helium or argon.3 For CAD, the precursor ion is accelerated to a certain kinetic energy, which is partially transferred to internal energy upon the collision. This internal energy is averaged out over all degrees of freedom of the molecule. If the energy supplied to a particular bond is higher than the bond energy, the bond breaks and forms a fragment.

This fragmentation can occur multiple times, until the energy remaining in the molecule is not sufficient to break any bonds. The maximum internal energy that the precursor ion can gain in a collision (Ecm) is related to the kinetic energy of the ion (Elab), the mass of

3 the collision gas (Mg) and the mass of the precursor ion (Mi). (Equation 2.2)

Mi Ecm = Elab Equation 2.2 Mg+Mi

Ecm and Elab are also known as center-of-mass and laboratory-frame kinetic energies of the precursor ion, respectively. Ecm is used to derive survival yield (SY) curves, as seen in Chapter IV.

2.5.3 Types of fragmentation

Depending on how the electrons move during bond clevages, two different types of fragments can be formed. These are illustrated in Scheme 2.1 for a positive precursor ion.

Scheme 2.1. Two different fragmentation mechanisms.

20 The first mechanism represents a homolytic bond cleavage and gives rise to radicals. The fragment that retains the charge will be the one seen in the fragmentation spectrum. The second mechanism represents a heterolytic bond cleavage. Again, the fragment retaining the positive charge is observed in the spectrum. In both cases, the charge remains preferentially on the fragment that leads to the thermodynamically most stable product.

2.6 Ion Mobility Mass Spectrometry (IM-MS)

Standard mass spectrometry separates ions by their mass and charge, but cannot distinguish isomers. This problem can be overcome by coupling ion mobility spectrometry to mass spectrometry. Ion mobility spectrometry was introduced in the late

1800’s with the X-ray experiments of Thomson and Rutherford, with hybrid ion mobility mass spectrometry instruments being developed in the 1960s.24,25 Ion mobility spectrometry really began to increase in popularity when it was discovered that the hyphenated ion mobility mass spectrometry (IM-MS) method could be used for structure- based characterization and differentiation of chemical isomers.26–29 The last two decades have seen major improvements in the coupling of ion mobility and mass spectrometry, including the invention of many different types of ion mobility including drift time ion mobility spectrometry, temporally-dispersive ion mobility spectrometry, overtone mobility spectrometry, and travelling wave ion mobility spectrometry (TWIMS), which is the type of ion mobility used in this dissertation.30

21 2.6.1 Travelling wave ion mobility spectrometry (TWIMS)

The traveling wave ion mobility technique was invented by Giles in 2004,31 and released commercially by Waters with their Synapt HDMS,32 the instrument used in this dissertation (Figure 2.10).

Figure 2.10. Traveling wave ion mobility spectrometry.

Reproduced from article (ACS does not require permission for individual figures).30

The ion mobility region consists of three sections: the trap cell, ion mobility cell, and transfer cell (Figure 2.11).

Figure 2.11. Triwave portion of the Synapt HDMS mass spectrometer.

Reproduced with permission.33

The trap cell is used to accumulate packets of ions before introducing them simultaneously into the ion mobility cell. In the ion mobility cell, electric pulses are used to move the ions along the cell, while a buffer gas flows in the opposite direction.31 This separates the ions by their size and shape. The transfer cell then keeps the separation of the ions while transferring them to the ToF mass analyzer. Due to the presence of trap and transfer cells, CAD can be performed either before or after ion mobility separation.34,35 Fragmentation in the trap cell followed by ion mobility analysis would separate the fragments. Fragmentation in the transfer cell would fragment ions after their separation by ion mobility.

By superimposing a direct current potential (pulse) on the ring electrodes in the ion mobility cell and switching that potential to an adjacent electrode, a moving electric

23 field, or ‘travelling wave’, is created on which the ions can ‘surf’.31 Ions with smaller mobilities will not collide with the buffer gas as frequently as ions with larger mobilities and will pass through the ion mobility cell more quickly. The height and velocity of travelling wave can be changed to improve the separation of the ions.

2.6.2 Collision Cross Sections (CCS)

Collision cross sections can be calculated from the drift time output of TWIMS experiements. With TWIMS, calibrants with known CCS values must be used to convert the measured drift time of an ion to a CCS. The equations used to derive the calibration equation are seen below (Equations 2.3 – 2.5).36 The drift time of the calibrant ions

(DTcal) are measured and corrected as shown in Equation 2.3. The known CCS values of the calibrant ions (Ωcal) are converted to mass- and charge-independent values as shown in Equations 2.4 and 2.5. Corrected CCS (Ω’) is plotted against corrected drift time

(DT’) to obtain the calibration curve and calibration equation (Equation 2.6).

1.4∗ 푀 퐷푇′ = 퐷푇 − √ 푖 Equation 2.3 푐푎푙 1000

푀 푀 휇 = 푔 푖 Equation 2.4 푀푔+푀푖

Ω ∗ 휇 Ω′ = 푐푎푙 √ Equation 2.5 푧

Ω′ = 퐴(퐷푇′)퐵 Equation 2.6

Variables in Equations 2.3 – 2.6: DT’ = corrected drift time, DTcal = drift time of the calibrant, Mi = mass of the ion, Mg = mass of the buffer gas in the ion mobility cell, Ω’ = corrected CCS, Ωcal = CCS of calibrant, μ = reduced mass of the ion/drift gas complex, A

= fitting parameter, B = fitting parameter, z = charge of ion.

24 Equation 2.6 is used to calculate the Ω’ of an unknown compound based on the measured DT’ of this compound. The following variables are necessary to calculate the

CCS of the unknown: Mi, DTunknown, and z. The CCS (or Ω) of the unknown can then be determined using Equation 2.7.

Ω′∗푧 Ω = Equation 2.7 √휇

CCS values derived this way are used to compare different oligomers or isomers of the same compound or compare and contrast compounds.

CHAPTER III

MATERIALS AND INSTRUMENTATION

3.1 Materials

Tetrahydrofuran (THF), methanol (MeOH), water (H2O), and acetonitrile (ACN) were purchased from Fisher Scientific (Waltham, MA), at Optima grade. The following were purchased from Sigma Aldrich (St. Louis, MO): dimethyl sulfoxide (DMSO,

Chromasolv, ≥99.7%), sodium trifluoroacetate (NaTFA, ≥99.0%), silver trifluoroacetate

(AgTFA, ≥99.99%), trans-2-[3-(4-tert-Butylphenyl)-2-methyl-2- propenylidene]malononitrile (DCTB, ≥99.0%), formic acid (~98%), and polyalanine

(molecular weight 1,000 – 5,000).

The copolyesters to be discussed in Chapter IV were synthesized by Mayela

Cristina Ramirez-Huerta, in Prof. Mark D. Soucek’s group at the Department of Polymer

Engineering of the University of Akron.37,38 The hyperbranched fluorinated polymers to be discussed in Chapter V were synthesized by Matthew Quast, in Prof. Anja Mueller’s group at Central Michigan University.39 The sugar-based isomers to be discussed in

Chapter VI were synthesized by Cesar Ganzalez, in Prof. Coleen Pugh’s group at the

Department of Polymer Science of the University of Akron.

All materials were used in the condition received from their supplier without further purification or modification.

26 3.2 Instrumentation

The following sections describe the mass spectrometers used in this dissertation, as well as the instrument setting used to obtain the data presented.

3.2.1 Synapt HDMS

The Synapt HDMS (Waters, Milford, MA) is an ESI-Q/ToF mass spectrometer equipped with a TWIMS cell and microchannel plate detector. (An explanation of these terms can be found in Chapter 2.) A scheme of this instrument can be seen in Figure 3.1.

Figure 3.1. Schematic diagram of the Synapt HDMS.

Reproduced with permission.40

The flow rate of the analyte into the instrument was kept constant at 10 uL/min. The capillary voltages was set to 3.16 kV, with the sampling cone and extraction cone

27 voltages set to 35 V and 3.2 V, respectively. The source temperature and desolvation gas temperature were set to 80°C and 150°C, respectively, while the desolvation gas flow rate was set at 500 L/h. During standard MS analysis, the trap and transfer potentials were set to 6.0 V and 4.0 V, respectively. The trap potential was changed for MS/MS experiments, as will be explained in each chapter. The trap gas (Ar) flow rate was kept constant at 1.5 mL/min. The ion mobility wave velocities, wave heights, and gas flow rates will be discussed in each chapter as necessary. The data were acquired using

MassLynx (version 4.1) and the ion mobility data analyzed using DriftScope (version

2.1).

3.2.2 Bruker Ultraflex III

The Bruker Ultraflex III (Bruker Daltonics, Billerica, MA) is a MALDI-ToF/ToF mass spectrometer equipped with a microchannel plate detector. The laser for the instrument is a Nd:YAG laser that emits light at a wavelength of 355 nm. The laser power of the instrument was adjusted for each project in order to maximize the signal intensity without causing fragmentation. The laser power will be discussed in each individual chapter. The ions were accelerated by 25 kV in the IS1 region. The IS2 voltage was set to 21.65 kV, the lens voltage to 9.65 kV, and the delay time was 150 ns.

The reflectron lenses were set at 26.30 and 17.30 kV, respectively.

There are two possible modes in which to operate the ToF/ToF, linear and reflectron mode. In this dissertation, all spectra were collected in reflectron mode. The instrument also contains a LIFT cell, which can be used for MS/MS. This portion of the

28 instrument was not used in this dissertation. The Ultraflex III uses flexControl (version

3.4) for data acquisition and flexAnalysis (version 3.4) for data analysis.

29 CHAPTER IV

POLYESTERS

4.1 Introduction

Aliphatic polyesters are biodegradable and biocompatible, which presents some unique applications. Their use in biodegradable fibers, films, and plastics has received significant attention in recent years.41 Their biodegradation is related to the polymerization reaction. A diol and diacid react to form the polymer, and the reverse reaction can occur in order for the polymer to biodegrade (Scheme 4.1).

Scheme 4.1. The condensation/hydrolysis reaction.

X is the R group of the diol. Y is the R group of the diacid.

In order to control the biodegradation, an understanding of the hydrolysis of the polyesters is necessary. The purpose of synthesizing these polymers was to study the influence of the backbone (i.e. diacid and diol) on the hydrolysis of the polyesters. To solely study that effect, the polymers were end-capped with urethane groups using phenyl isocyanate, in order to prevent the hydroxyl end groups from causing the degradation reaction by transesterification. Five different copolyesters were synthesized (Scheme

4.2).

30 O O AA.NPG O AA.1,2-EG O O O NH HN O O HN O O O O NH n O O O O n

O O CHDA.NPG O CHDA.1,2-EG HN O HN O O O O O O NH O O O NH n O O O n

O O CHDA.1,5-PED

HN O O O O NH

O n O

Scheme 4.2. The five copolyesters to be discussed in this Section.

The polymers investigated were synthesized using two different diacids and three different diols. The two diacids were 1,4-cyclohexane dicarboxylic acid (CHDA) and adipic acid (AA). The three diols were 1,5-pentane diol (1,5-PED), 1,2-ethylene glycol

(1,2-EG), and neopentyl glycol (NPG).

Polyesters have been well characterized by mass spectrometry.42–50

When energetically activated, polyester ions dissociate largely by 1,5-hydrogen rearrangement as seen in Scheme 4.3.42,51–53 This resembles the McLafferty rearrangement occurring in radical ions54 and is also observed upon the thermal degradation of polyesters.55,56

Scheme 4.3. The 1,5-hydrogen rearrangement,

as it applies to copolyesters.

As can be seen, the 1,5-hydrogen rearrangement requires the presence of a hydrogen atom on the β-carbon of the diol. A hydrogen atom on the β-carbon is present on the 1,2-

EG and 1,5-PED diols, but is absent from the NPG diol. The three copolyesters containing the 1,2-EG and 1,5-PED diols can thus fragment via a 1,5-hydrogen rearrangement, but the two copolyesters containing the NPG diol cannot. Interestingly, the isomeric copolyesters CHDA.NPG and CHDA.1,5-PED were found to undergo very similar fragmentations, leading to indistinguishable MS/MS spectra (vide infra). This requires proposing different fragmentation mechanism for these two copolyesters.

Mass spectrometry is a powerful analysis technique; however, single-stage mass spectrometry is not capable of distinguishing isomers. In response to this weakness, complimentary techniques are required.57–61 Generally, liquid chromatography can be used to separate isomers of different polarities,59 and tandem mass spectrometry

(MS/MS) can be used to distinguish isomers based on their fragmentation pathways.57–62

Ion mobility mass spectrometry (IM-MS) and collision cross sections (CCS) are used to distinguish isomers as well.63–70 MS/MS, IM-MS, and CCSs will be compared for their ability to separate and distinguish the two structural isomers: CHDA.NPG and

CHDA.1,5-PED.

32 This chapter is divided into two sections. Section 4.3.1 focuses on the analysis of the two structural isomers, CHDA.1,5-PED and CHDA.NPG. It discusses the use of

MS/MS, IM-MS, and CCS data to characterize and distinguish the two isomers. This section also describes a new fragmentation mechanism for CHDA.NPG, which cannot undergo the 1,5-hydrogen rearrangement. Section 4.3.2 focuses on the analysis of the three additional copolyesters, as well as on comparing and contrasting the results of all five copolyesters.

4.2 Experimental Methods

The following sections discuss the sample preparation methods and experimental parameters used in the rest of the chapter.

4.2.1 Synthesis of Polyesters

Five different polyesters were synthesized by Mayela Cristina Ramirez-Huerta in

Prof. Mark D. Soucek’s research group at the Department of Polymer Engineering of the

University of Akron.37,38 A molar excess of the diol was used to produce bis-hydroxy- terminated chains. The reaction proceeded under nitrogen and was catalyzed with dibutyltin oxide. The temperature of the reaction was varied as a function of time to optimize the reaction. The polyesters formed this way were dried under vacuum for 5 hours at 110°C. The hydroxyl-terminated polyesters were end-capped using phenyl isocyanate by adding a 1.1:1 molar excess of the phenyl isocyanate and using dibutyltin dilaurate as a catalyst. IR was used to monitor the reaction in order to study the change

33 in intensity of the hydroxyl peak over time. Once the hydroxyl peak in the IR spectrum had disappeared, the reaction was considered complete.

The polymers were hydrolyzed in acetone with a large excess of water. The five polyesters were each placed in the hydrolysis solution and allowed to undergo hydroysis for different times as described in Table 4.1.

Table 4.1. Hydrolysis times of the various polyesters.

Hydrolysis Times (days) CHDA.1,5-PED 13 51 85 CHDA.1,2-EG 4 34 82 CHDA.NPG 19 45 74 AA.1,2-EG 7 28 82 AA.NPG 68 96

When the times listed were reached, an aliquot of the solution was removed and dried in a convection oven at 100°C for three hours. These samples were submitted for mass spectrometry analysis.

4.2.2 MALDI Preparation

Each of the polyesters and the salt (NaTFA) were dissolved at a concentration of

10 mg/mL in THF. The matrix, DCTB, was dissolved at a concentration of 20 mg/mL in

THF. The matrix, sample, and salt were combined at a ratio of 100:20:10 (v/v). A droplet of the mixture (approximately 0.6 uL) was added to the MALDI plate drop-wise and allowed to dry before analysis.

34 4.2.3 ESI Preparation

Each polyester sample was dissolved in 9:1 (v:v) THF:MeOH at a concentration of 0.1 mg/mL. One uL of NaTFA solution (at a concentration of 10 mg/mL in THF) was added to 1000 uL of the sample solution. Ubiquitin (as a standard for IM-MS) was dissolved in 5:5 H2O:ACN at a concentration of 0.68 mg/mL. Twenty uL of acetic acid was added to 1000 uL of the ubiquitin solution.

4.2.4 Synapt Q/ToF Parameters

The trap potential was varied from 10 – 90 V, corresponding to a collision energy

(CE) window of 10 – 90 eV for the survival yield curves. (Additional information is given in Section 4.2.5.) The IM-MS parameters were as follows: travelling wave height

7.0 V, travelling wave velocity 200 m/s, and bath gas flow rate 22.7 mL/min.

4.2.5 Survival Yield Calculations

In order to construct the survival yield curves, several n-mers (either 1-mer to 3- mer or 1-mer to 4-mer) from each of the five polyesters was fragmented via CAD over a range of collision energy values. The collision energy ranges used can be seen in Table

4.2. In general, data were collected every 10 eV, with the interval being reduced to 5 eV closer to the E50 value. The entire survival yield data were collected in triplicate.

35 Table 4.2. The collision energies used for each oligomer of the five copolyesters.

1-mer 2-mer 3-mer 4-mer (eV) (eV) (eV) (eV) CHDA.1,5-PED 10 to 50 10 to 65 10 to 90 CHDA.1,2-EG 10 to 40 10 to 50 10 to 60 10 to 70 CHDA.NPG 10 to 60 10 to 75 10 to 90 AA.1,2-EG 10 to 35 10 to 40 10 to 50 10 to 60 AA.NPG 10 to 50 10 to 65 10 to 80

After the CAD spectra of an individual oligomer were collected, survival yields were calculated for each CE by comparing the intensities of the peaks, as seen in

Equation 4.1, where IP is the intensity of the parent ion and ΣIF is the sum of the intensity of all fragments.71

I SY = P Equation 4.1 IP+∑ IF

This equation will give a survival yield between zero and one. This value is then compared to the corresponding Ecm (as explained in Section 2.5.2).

The SY is plotted against Ecm to make a survival yield curve graph. Such graphs have sigmoid shape, similar to the shape of titration curves.71 The equation of the resulting sigmoidal line was calculated using Igor Pro, which gave the coefficients and error of the coefficients for the sigmoidal curve, as seen in Equation 4.2.

(B±b) 푆Y = (A ± a) + −(x−(C±c)) 1+e (D±d)

Equation 4.2

The coefficients calculated for each line are A, B, C, and D, and their respective errors

71 are a, b, c, and d. An important value derived from survival yields is the E50 value.

This is the Ecm at which the survival yield is equal to 0.5 or 50%. This is a characteristic

36 fragmentation energy for each of the examined n-mers, and can be used to compare and

72 contrast the samples. In order to calculate the E50 value for an individual n-mer, the coefficients and their errors calculated from the corresponding sigmoidal curve were taken into account, as shown in Equation 4.3. The derivation of the equation is provided in Appendix II.

2 2 퐵 푏 2 푎 2 √( ) +( ) 0.5−퐴 퐵 0.5−퐴 퐵 2 −1 퐵 2 퐵 푑 0.5−퐴 퐸50 = ((−퐷) ln ( − 1) + 퐶) ± 푐 + (−퐷) ln ( − 1) ( ) + 퐵 0.5−퐴 0.5−퐴 퐷 ln( −1) 0.5−퐴

√ ( ) √ ( )

Equation 4.3

The first part the equation reveals the E50 value, and the square root term calculates the error. These equations were used to calculate the survival yield curves and E50 values discussed in Sections 4.3.1 and 4.3.2.

4.3 Results

The following two sections discuss the analysis of CHDA.NPG, CHDA.1,5-PED,

CHDA.1,2-EG, AA.NPG, and AA.1,2-EG.

4.3.1 Analysis of CHDA.NPG and CHDA.1,5-PED

The MALDI spectra of CHDA.NPG (Figure 4.1) and CHDA.1,5-PED (Figure

4.2) can be seen below.

37 Figure 4.1. MALDI spectra of CHDA.NPG

after three hydrolysis times.

38 Figure 4.2. MALDI spectra of CHDA.1,5-PED

after three hydrolysis times.

As would be expected for structural isomers, the masses of the oligomers in the two

Figures are identical within experimental error. There are differences in the relative abundances between the two polyesters, indicating differences in hydrolysis behavior.

This aspect will be discussed together with all other polyesters in section 4.3.2.

The MS/MS spectrum of the 3-mer from CHDA.1,5-PED, acquired via ESI-CAD can be seen in Figure 4.3. CHDA.1,5-PED fragments mainly via 1,5-H rearrangements, rationalized in Scheme 4.4. In addition, phenylisocyanate is lost from the urethane end groups (Scheme 4.5).

39 Figure 4.3. MS/MS spectrum of the sodiated 3-mer (1085 m/z) of CHDA.1,5-PED.

Monoisotopic m/z values are given on select peaks. The m/z values of the fragments

formed by consecutive dissociations are underlined.

Scheme 4.4. 1,5-Hydrogen rearrangement in collisionally activated CHDA.1,5-PED.

40 Scheme 4.5. Phenyl isocyanate loss from the copolyester end groups.

Loss of one end group (Scheme 4.5) and random 1,5-H rearrangement fragmentations over the ester groups in the polymer chain (Scheme 4.4) lead to the major fragments at m/z 966, 948, 880, 708, 640, 468, and 400, as summarized in Scheme 4.6. Consecutive fragmentation via the same mechanisms accounts for all other, less abundant fragments.

All fragments formed by these dissociations are linear, as denoted by the ‘l’ in the fragment identification. The subscript is the number of repeat units present in the fragment. The end groups of the fragment are denoted as superscripts (Scheme 4.7), with

H denoting a hydroxyl end group (from the diol), F denoting phenyl isocyanate, V denoting a vinyl (alkene) end group (from the diol), and A denoting an acid end group

(from the diacid).

While the CHDA.1,5-PED fragmentation pathways are easily explained via 1,5- hydrogen rearrangements, CHDA.NPG cannot undergo this reaction. However, identical fragments are seen in the CHDA.NPG MS/MS spectrum, (Figure 4.4) albeit with somewhat different relative abundances, especially below 640 m/z. This means another fragmentation mechanism must operate for CHDA.NPG, which prompted additional experiments to elucidate the seemingly similar fragmentation pattern.

41 spectrum of sodiated CHDA.1,5- PED trimer. Scheme 4.6. Major and minor fragments in the MS/MS

42 Scheme 4.7. CHDA.1,5-PED fragment end group structures.

Figure 4.4. MS/MS spectra of the sodiated 3-mer of CHDA.NPG (1085 m/z).

Monoisotopic m/z values are given of select peaks. The m/z values of the fragments

formed by consecutive dissociations are underlined.

43 More information on the fragmentation mechanism of CHDA.NPG was sought by comparing its fragmentation energetics to those of CHDA.1,5-PED. For this, survival yields (SY) were measured at different collision energies and plotted against the corresponding center-of-mass energies (Ecm) as described above. From the survival yield curves constructed this way, E50 values can be calculated, which as explained in Section

2.5.2, represent fragmentation energies for the dissociating oligomers.

SY measurements were repeated in triplicate, and the average data sets used to construct the SY curves (Figures 4.5 – 4.6) and calculate the E50 values. The coefficients and errors for the sigmoidal curves can be seen in Table 4.3. (The equation of the curve using those parameters and errors is given in Section 4.2.5, Equation 4.2.) The

71 characteristic E50 values and their error, calculated from those curves, are included in

Table 4.4.

Table 4.3. Values of the coefficients and errors for the survival yield curves for

CHDA.NPG and CHDA.1,5-PED.

A B C D a b c d CHDA.NPG 1-mer 1.0127 -1.0559 1.4149 0.1875 0.0052 0.0127 0.0054 0.0049 CHDA.NPG 2-mer 0.9853 -0.9824 2.2554 0.2294 0.0042 0.0080 0.0064 0.0058 CHDA.NPG 3-mer 0.9707 -0.9623 2.2060 0.1879 0.0060 0.0131 0.0089 0.0077

CHDA.1,5-PED 1-mer 1.0004 -1.2134 1.5930 0.2172 0.0099 0.1029 0.0382 0.0178 CHDA.1,5-PED 2-mer 0.9876 -0.9764 2.3459 0.2540 0.0039 0.0110 0.0075 0.0064 CHDA.1,5-PED 3-mer 0.9434 -0.9334 2.2190 0.2229 0.0046 0.0116 0.0080 0.0071

44 CHDA.NPG 1 0.9 0.8

0.7

1-mer 0.6 2-mer 0.5 3-mer 0.4

1-mer Data Survival Yield 0.3 2-mer Data 0.2 3-mer Data 0.1 0 0.00 1.00 2.00 3.00 4.00 Energy of Center of Mass (eV)

Figure 4.5. Survival yield curves for sodiated CHDA.NPG oligomers.

CHDA.1,5-PED 1 0.9 0.8

0.7

1-mer 0.6 2-mer 0.5 3-mer 0.4

1-mer Data Survival Yield 0.3 2-mer Data 0.2 3-mer Data 0.1 0 0.00 1.00 2.00 3.00 4.00 Energy of Center of Mass (eV)

Figure 4.6. Survival yield curves for sodiated CHDA.1,5-PED oligomers.

45 Table 4.4. E50 values for CHDA.NPG and CHDA.1,5-PED oligomers

CHDA.NPG CHDA.1,5-PED (eV) (eV) 1-mer 1.404±0.007 1.516±0.050 2-mer 2.250±0.008 2.345±0.010 3-mer 2.198±0.010 2.197±0.010

The E50 values for CHDA.NPG and CHDA.1,5-PED show certain trends. The values are significantly different for the 1-mers, more similar but still distinguishable for the 2-mers, and indistinguishable for the 3-mers. The different E50 values for the 1-mer and 2-mer are consistent with different fragmentation mechanisms for CHDA.NPG and CHDA.1,5-

PED. The progressive similarity of the E50 values as the polyester chains become longer further suggests that the fragmentation of CHDA.NPG has similar energetics to the

McLafferty rearrangement in CHDA.1,5-PED.

In order to determine whether size or architectural differences affect the fragmentation mechanism, CCSs were deduced for several oligomers of both polyesters, via ion mobility mass spectrometry (IM-MS). Figures 4.7 and 4.8 show the IM-MS plots of CHDA.1,5-PED and CHDA.NPG respectively. These plots indicate the drift times of the various polyester n-mers through the IM region. In order to calculate CCS values from the drift times, the drift time scale must be calibrated with standards of known CCS, as mentioned in the Experimental Section. The standards used were different charge states of ubiquitin, which was analyzed under the same IM-MS conditions as CHDA.1,5-

PED and CHDA.NPG. Table 4.5 lists the measured drift times of the calibrant ions as well as their reported CCSs and the corrected drift time and CCS data calculated by the

46 procedure outlined in Section 2.6.2. A plot of Ω’ vs DT’ gives the calibration curve in

Figure 4.9.

Figure 4.7. IM-MS plot (m/z vs. drift time) of CHDA.1,5-PED. The [M+Na]+ ions of

intact oligomers are encased. The number of repeat units is also given.

Figure 4.8. IM-MS plot (m/z vs. drift time) of CHDA.NPG. The [M+Na]+ ions of intact

oligomers are encased. The number of repeat units is also given.

Table 4.5. Calibration Data

DT(a) DT'(b) Ω (CCS) Ω' (CCS z m/z μ (ms) (ms) (Å2)(c),73 norm.) (Å2)(d)

4 2140.3 5.05 4.99 1004 5.2582 1319.80 5 1712.2 4.06 4.00 1137 5.2497 1193.78 6 1426.8 5.41 5.36 1525 5.2413 1332.15 7 1223.0 4.42 4.37 1580 5.2329 1181.13 8 1070.1 3.34 3.29 1622 5.2245 1059.27 9 951.2 2.71 2.67 1649 5.2162 955.72 10 856.1 2.35 2.31 1732 5.2079 902.01 11 778.3 2.17 2.13 1802 5.1997 851.81

(a) Measured drift time.

(b) Corrected drift time (Equation 2.3).

(d) Normalized CCS (“charge and mass independent” CCS; Equation 2.5)

y = 595.38x0.4847 Ubiquitin R² = 0.9903 1600

1400

) 1200 2 Å 1000 800 600

Norm. Norm. CCS( 400 200 0 1.5 2.5 3.5 4.5 5.5 6.5 Corr. DT' (ms)

Figure 4.9. Calibration curve for ubiquitin.

48 The equation of the calibration curve can be used with the drift times for CHDA.NPG and CHDA.1,5-PED to determine their experimental CCSs, again using the equations found in Section 2.6.2. The IM-MS measurements were repeated in triplicate, so that standard deviations could also be calculated (Table 4.6).

Table 4.6. Experimental CCS for CHDA.NPG and CHDA.1,5-PED

CHDA.NPG CHDA.1,5-PED DT(a) DT'(b) Ω'(c) Ω(d) DT(a) DT'(b) Ω'(c) Ω(d) [M+Na]+ m/z (ms) (ms) (Å2) (Å2) (ms) (ms) (Å2) (Å2) 2.62 2.59 944 2.35 2.32 895 1-mer 605 2.71 2.68 959 185±1 2.44 2.41 911 175±1 2.71 2.68 959 2.44 2.41 911 4.51 4.47 1230 4.33 4.29 1206 2-mer 845 4.51 4.47 1230 237±1 4.42 4.38 1218 234±2 4.6 4.56 1242 4.51 4.47 1230 6.41 6.36 1460 6.32 6.27 1450 3-mer 1085 6.5 6.45 1470 281±2 6.41 6.36 1460 279±1 6.59 6.54 1480 6.5 6.45 1470 8.39 8.34 1664 8.5 8.45 1675 4-mer 1325 8.48 8.43 1673 320±1 8.57 8.52 1682 321±1 8.57 8.52 1682 8.66 8.61 1690

(a) Measured drift time. (See Figures 4.7 and 4.8)

(b) Corrected drift time (Equation 2.3).

(d) Experimental CCS (Equation 2.7).

A small difference in compactness is observed for the 1-mer and an even smaller one for the 2-mer. All other oligomers have essentially identical CCSs and sizes. This suggests similar size effects on the fragmentation mechanisms of CHDA.NPG and CHDA.1,5-

PED.

49 Theoretical values for the CCS were also calculated and compared to the experimental values. Fifty structures were modeled by molecular mechanics/dynamics simulations using the Materials Studio software three different times for a total of 150 structures. The coordinates of the energy-minimized structures were then input into the

MOBCAL program to determine the corresponding CCS. A plot of calculated CCS vs. relative energy of the 150 calculated structures for the CHDA.1,5-PED 2-mer is shown in

Figure 4.10. A representative structure is depicted in Figure 4.11, together with a representative structure of CHDA.NPG to illustrate the corresponding architectures.

CHDA.1,5-PED 2-mer 300

275

250

225 PA (CCS) 200

175

150 150 175 200 225 250 275 300 Energy

Figure 4.10. Calculated CCS vs. relative energy for 150 energy-minimized structures of

the CHDA.1,5-PED 2-mer. The three different sets of 50 are designated by the three

different colors.

50 The theoretical CCSs can be seen in Table 4.7. When comparing the theoretical and experimental values, there is <2% discrepancy, with the exception of the CHDA.1,5-PED

1-mer for which measured and calculated CCS agree within ~7%. Agreement within

~10% is generally satisfactory to conclude that the calculated architectures reconcile with the measured drift times and CCSs.

Table 4.7. Theoretical CCS for CHDA.NPG and CHDA.1,5-PED

CCS (Å2) (a) CHDA.NPG CHDA.1,5-PED 1-mer 175.56±2.70 176.02±7.64 2-mer 224.27±8.99 227.81±13.98 3-mer 264.17±13.19 269.67±18.58 4-mer 303.22±18.62 301.02±22.25

(a) Calculated from simulated structures using the projection approximation method in the

MOBCAL algorithm (mean and std. deviation of the 150 values).

51 Figure 4.11. Representative energy-minimized structures for the sodiated 2-mers from

(a) CHDA.NPG and (b) CHDA.1,5-PED

52 Based on the SY and CCS data provided above, the charge-induced mechanism in

Scheme 4.8 is proposed for the MS/MS decomposition of sodiated CHDA.NPG.

Scheme 4.8. Proposed fragmentation mechanism for the COO-CH2 bond cleavage in

sodiated CHDA.NPG

+ Na coordination at the ester carbonyl group weakens the neighboring CH2-O(CO) bond, facilitating heterolytic bond cleavage. The recipient product of such dissociation would be a primary carbenium ion, which is highly unstable. The NPG connectivity allows for a

1,2-CH3 shift, however, yielding a much more stable tertiary carbocation on one fragment and a sodium carboxylate ion pair (salt bridge) on the other. The negatively charged carboxylate ion abstracts a proton from the carbocation to form a double bond. This fragmentation mechanism would produce fragments with identical m/z values as those resulting by 1,5-hydrogen rearrangement from sodiated CHDA.1,5-PED, as can be seen in the MS/MS spectra of Figures 4.3 and 4.4. The SY data point out that the charge- induced fragmentation pathway shown in Scheme 4.8 is energetically more favorable than the 1,5-H rearrangement pathway shown in Scheme 4.4 for oligomers less than 3 repeat units (vide supra). As the chain length increases, the energy requirements of these

53 two pathways become indistinguishable. The superscripts used to designate the end groups of the fragments generated by one or successive dissociations according to the mechanism of Scheme 4.8 are listed in Scheme 4.9.

Scheme 4.9. CHDA.NPG fragment end group structures.

In addition to elucidating the fragmentation mechanism of CHDA.NPG, a goal of this dissertation was to determine a means to distinguish the CHDA.NPG and CHDA.1,5-

PED isomers using mass spectrometry. Because the mass spectra and tandem mass spectra match, the SY curves and IM-MS data need to be used for this purpose. Using either the E50 values or CCSs, the lower molecular weight oligomers (i.e. the 1-mer and

2-mer) can be differentiated. However, as the molecular weight of the oligomers increases, the isomers become indistinguishable by MS methods. If MS must be used, because the sample is a blend or not available in pure form, the low molecular weight oligomers should be used to determine which of the isomers is present.

54 4.3.2 Analysis of AA.1,2-EG, CHDA.1,2-EG, and AA.NPG

This section will discuss the analysis of the other three copolyesters (AA.1,2-EG,

CHDA.1,2-EG, and AA.NPG) as well as draw general conclusions about all five copolyesters. The MALDI spectra of AA.1,2-EG (Figure 4.12), CHDA.1,2-EG (Figure

4.13), and AA.NPG (Figure 4.14) agree well within their expected repeat units and end groups.

Figure 4.12. MALDI mass spectra of AA.1,2-EG after three hydrolysis times.

Figure 4.13. MALDI mass spectra of CHDA.1,2-EG after three hydrolysis times.

56 Figure 4.14. MALDI mass spectra of AA.NPG after two hydrolysis times.

The Mn values of the hydrolysis products from each of the five copolyesters were calculated from the corresponding MALDI mass spectra using the Polymerix software available in our group. If the samples were being hydrolyzed and degraded, the Mn values would be expected to shift down. This is the case for AA.1,2-EG, and to some extent, for CHDA.1,5-PED. However, the other three copolyesters show Mn values that increase (AA.NPG and CHDA.1,2-EG) or increase and then decrease (CHDA.NPG) with hydrolysis time. An increase in Mn could arise if shorter oligomers depolymerize fast, while the longer chains remain largely intact. In cannot be excluded, however, that there

57 was an error in the hydrolysis methodology. The hydrolysis experiments are currently being repeated in the Soucek group for a definitive answer.

Table 4.8. Mn values of the hydrolysis products of the five copolyesters.

Hydrolysis Time Mn (Days) AA.1,2-EG 7 2151 28 1528 82 1352 AA.NPG 68 1911 96 1942 CHDA.1,2-EG 4 2144 45 2225 84 2271 CHDA.1,5-PED 13 2392 51 2392 85 2307 CHDA.NPG 19 2099 45 2405 74 2208

The MS/MS fragmentation pathways of AA.1,2-EG and CHDA.1,2-EG were straightforward, since both polyesters can undergo the 1,5-hydrogen rearrangement. The

MS/MS spectrum of CHDA.1,2-EG can be seen in Figure 4.15, and that of AA.1,2-EG in

Figure 4.16. The nomenclature for the fragments from AA.1,2-EG matches the one used for CHDA.1,2-EG, which is explained below.

58 Figure 4.15. The MS/MS spectrum of the sodiated CHDA.1,2-EG 4-mer

acquired at a CE of 55 eV.

As with CHDA.NPG and CHDA.1,5-PED, all fragments are linear, which is denoted by the ‘l’. The subscript identifies the number of repeat units, whereas the superscripts identify the end groups. The end groups are symbolized as follows: H denoting a hydroxyl end group (from the diol), F denoting phenyl isocyanate, V denoting a vinyl

(alkene) end group (from the diol), and A denoting an acid end group (from the diacid).

(Scheme 4.7, with 1,2-EG replacing 1,5-PED)

Figure 4.16. The MS/MS spectrum of the sodiated AA.1,2-EG 4mer

acquired at a CE of 45 eV.

The fragmentation behavior of AA.NPG is more complicated, since it cannot undergo the 1,5-hydrogen rearrangement. The polymer is assumed to dissociate via the same mechanism as CHDA.NPG, cf. Scheme 4.8. Other than formed through a different mechanism, the fragments of AA.NPG are similar to CHDA.1,2-EG and AA.1,2-EG, as attested by an inspection of the MS/MS spectra in Figures 4.15, 4.16, and 4.17. The end group classifications are the same as explained above (see also Scheme 4.9).

60 Figure 4.17. The MS/MS spectrum of the sodiated AA.NPG 3-mer

acquired at a CE of 65 eV.

For a better understanding of the energetics of the fragmentation, survival yield curves were measured for the three polymers. As explained in Section 4.2.5, [M+Na]+ ions of select oligomers were fragmented using a range of collision energies, dependent on the oligomer (Table 4.2). The survival yield curves obtained for the three polymers can be seen in Figures 4.18, 4.19, and 4.20 for AA.1,2-EG, CHDA.1,2-EG, and AA.NPG, respectively. The values for the coefficients and errors for curve-fitting the experimental data are given in Table 4.8 (The equation of the curve using those parameters and errors are described in Section 4.2.5.) The resulting E50 values and their uncertainties can be seen in Table 4.9. These data were calculated as explained in Section 4.2.5.

61 AA.1,2-EG 1

0.9

0.8

0.7 1-mer

2-mer 0.6 3-mer 0.5 4-mer 0.4 1-mer Data Survival Yield 0.3 2-mer Data 0.2 3-mer Data

0.1 4-mer Data

0 0.00 0.50 1.00 1.50 2.00 2.50 3.00 Energy of Center of Mass (eV)

Figure 4.18. Survival yield curves for sodiated AA.1,2-EG oligomers.

CHDA.1,2-EG 1 0.9 0.8 1-mer 0.7

2-mer 0.6 3-mer 0.5 4-mer 0.4 1-mer Data Survival Yield 0.3 2-mer Data 0.2 3-mer Data 0.1 4-mer Data 0 0.50 1.00 1.50 2.00 2.50 3.00 Energy of Center of Mass (eV)

Figure 4.19. Survival yield curves for sodiated CHDA.1,2-EG oligomers.

62 AA.NPG 1

0.9

0.8

0.7

1-mer 0.6 2-mer 0.5 3-mer 0.4 1-mer Data Survival Yield 0.3 2-mer Data 0.2 3-mer Data

0.1

0 0.00 1.00 2.00 3.00 4.00 Energy of Center of Mass (eV)

Figure 4.20. Survival yield curves for sodiated AA.NPG oligomers.

Table 4.9. Values of the coefficients and errors for the survival yield curves for

AA.1,2-EG, CHDA.1,2-EG, and AA.NPG.

Values of Coefficients and Errors A B C D a b c d AA.1,2-EG 1-mer 1.0027 -1.0045 1.5988 0.2093 0.0039 0.0058 0.0036 0.0034 AA.1,2-EG 2-mer 0.9779 -0.9836 1.5588 0.1740 0.0030 0.0050 0.0026 0.0024 AA.1,2-EG 3-mer 0.9539 -0.9617 1.5336 0.1574 0.0045 0.0076 0.0040 0.0039 AA.1,2-EG 4-mer 0.9759 -0.9813 1.4935 0.1470 0.0039 0.0062 0.0034 0.0032

CHDA.1,2-EG 1-mer 1.0046 -1.0085 1.7224 0.2228 0.0040 0.0060 0.0041 0.0039 CHDA.1,2-EG 2-mer 0.9471 -0.9544 1.6957 0.1879 0.0054 0.0086 0.0055 0.0049 CHDA.1,2-EG 3-mer 0.8696 -0.8736 1.6931 0.1621 0.0040 0.0067 0.0043 0.0038 CHDA.1,2-EG 4-mer 0.8590 -0.8737 1.6245 0.1755 0.0106 0.0188 0.0118 0.0111

AA.NPG 1-mer 0.9597 -0.9808 2.3169 0.2853 0.0062 0.0140 0.0103 0.0095 AA.NPG 2-mer 0.9634 -0.9632 2.1402 0.2126 0.0034 0.0061 0.0044 0.0039 AA.NPG 3-mer 0.8794 -0.8787 2.0894 0.1885 0.0057 0.0101 0.0076 0.0067

Table 4.10. E50 values for AA.1,2-EG, CHDA.1,2-EG, and AA.NPG oligomers.

AA.1,2-EG CHDA.1,2-EG AA.NPG (eV) (eV) (eV) 1-mer 1.599±0.005 1.723±0.005 2.281±0.013 2-mer 1.549±0.003 1.672±0.007 2.124±0.005 3-mer 1.516±0.005 1.643±0.005 2.038±0.009 4-mer 1.485±0.004 1.561±0.145

The survival yield curves of the three polymers follow the same trends, with the E50 values gradually decreasing with increasing molecular weight of the oligomers. This trend is different from the one found for CHDA.NPG and CHDA.1,5-PED. For both those polymers, the 1-mer had the lowest E50 value. However, above the 1-mer,

CHDA.NPG and CHDA.1,5-PED followed the same trend as the other three polymers, with higher molecular weight oligomers having lower E50 values.

Based on the survival yield curves and E50 values of the five copolyesters, the effect of the diacid and diol on the fragmentation energetics can be deciphered. Table

4.4, containing the E50 values of CHDA.NPG and CHDA.1,5-PED, is replicated here for comparison of the fragmentation energetics of all five copolyesters.

Table 4.4. E50 values for CHDA.NPG and CHDA.1,5-PED oligomers.

CHDA.NPG CHDA.1,5-PED (eV) (eV) 1-mer 1.404±0.007 1.516±0.050 2-mer 2.250±0.008 2.345±0.010 3-mer 2.198±0.010 2.197±0.010

64 By comparing CHDA.1,2-EG to AA.1,2-EG, it can be concluded that changing the diacid from AA to CHDA increases the energy required to fragment the polymer.

The trend is less obvious when comparing CHDA.NPG and AA.NPG, due to the irregular order of the E50 values of CHDA.NPG. However, if the E50 value of the CHDA.NPG

1-mer is neglected, the same trend is seen: chains with the CHDA diacid require more energy to fragment than chains containing the AA diacid. A possible explanation of this trend is that AA is more flexible than CHDA, which could result in less energy being required to fragment the AA-based polymers versus the CHDA-based polymers.

However, it must be said that the difference between the E50 values is quite small.

The effect of the diol on the fragmentation energetics can also be studied. When comparing AA.1,2-EG versus AA.NPG, it is obvious that the polyester with the 1,2-EG diol requires significantly less energy to fragment than the polyester with the NPG diol.

This is also seen when comparing the values of CHDA.1,2-EG to the values of

CHDA.NPG and CHDA.1,5-PED. The 1,2-EG diol leads to significantly lower energy requirements than the 1,5-PED diol and NPG diol (except for the 1-mer). As concluded in Section 4.3.1.3, the fragmentation energetics of 1,5-PED and NPG based polyesters are not significantly different.

Therefore the following conclusions can be made regarding the amount of energy necessary to fragment sodium cationized polymers based on the identity of the diacid and diol:

AA < CHDA

1,2-EG < 1,5-PED ≈ NPG

65 4.4 Conclusions

The first section of this chapter focused on the analysis of CHDA.NPG and

CHDA.1,5-PED. Two different topics were covered: determining the fragmentation mechanism for CHDA.NPG and using novel mass spectrometry techniques to differentiate the isomers. A fragmentation mechanism for CHDA.NPG was proposed as seen in Scheme 4.8. The E50 values and CCSs of the CHDA.NPG and CHDA.1,5-PED

1-mers and 2-mers showed differences significant enough for differentiation, but the higher molecular weight oligomers were indistinguishable.

The second section of the chapter covered the analysis of CHDA.1,2-EG,

AA.NPG, and AA.1,2-EG. In regards to how diacid and diol identity affected the fragmentation, the following conclusions were made, in order from least amount of energy required to most.

1,2-EG < 1,5-PED ≈ NPG

AA < CHDA

The hydrolysis degradation of all five copolyesters was also briefly assessed. No consistent trend was found, suggesting different hydrolysis mechanisms for polyesters with the diacids and diols examined, or significant errors in the hydrolysis procedure used.

66 CHAPTER V

HYPERBRANCHED FLUORINATED POLYMERS

5.1 Introduction

Fluorinated polymers have a chemical inertness and an oxidative and hydrolytic stability that provides an excellent barrier against chemical and environmental wear.74–77 These properties, especially their high hydrophobicity, complicate the mass spectrometry analysis. Electrospray ionization (ESI) in particular is complicated by the hydrophobicity of these materials, such that the standard sample preparation methods do not produce ions. Ion formation has been enabled by addition of dimethyl sulfoxide

(DMSO),78,79 acting in a similar way as the supercharging agents used in protein mass spectrometry.80–84

Hyperbranched polymers have lower solution viscosity and higher solubility than their linear counterparts and, hence, are more suitable for applications requiring such properties.85,86 The fluorinated polymer studied in this dissertation can be seen in

Scheme 5.1. Hyperbranched architectures are produced if both fluorinated benzyl rings react during the polymerization (vide infra). It is difficult to determine if the polymer has such a structure through conventional analytical techniques, such as UV-VIS, IR, or even

NMR spectroscopy, without the availability of standards. Here, mass spectrometry techniques will be used to determine the extent of hyperbranching. Particularly promising would be ion mobility mass spectrometry (IM-MS), which has proven useful

67 in distinguishing the structures of linear and cyclic materials;53,70,87–94 tandem mass spectrometry has also been used for the structure determination of polymers.53,56,62,68,95–103

5.2 Experimental Methods

The following sections discuss the sample preparation methods and experimental parameters used in the rest of the chapter.

5.2.1 Synthesis of the Fluorinated Polymer

The polymer was synthesized by Matthew Quast in the research group of Anja

Mueller at Central Michigan University.39 3,5-Bis[(pentafluorobenzyl)oxy]benzyl alcohol was polymerized in the presence K2CO3 and 18-crown-6 in a dry solvent at room temperature for 24 hours. (Scheme 5.1)

F F

F F F F F O F F F F F F O O F F F F F F O O O F F OH F F O F F O O F F F F F F OH O F F F F F

3,5-bis[(pentafluorobenzyl)oxy]benzyl alcohol F F

Scheme 5.1. Polymerization of Fluorinated Polymer.

5.2.2 MALDI Preparation

The polymer and silver trifluoroacetate (AgTFA) were dissolved in THF at a concentration of 10 mg/mL. DCTB was dissolved in THF at a concentration of 20 mg/mL. The matrix, polymer, and salt were combined at a ratio of 10:5:1 (v:v:v), respectively. The solution was added to the MALDI plate by the dry-drop method.

68 5.2.3 ESI Preparation

Due to the high fluorine content, the polymer is highly hydrophobic, making ESI analysis difficult. DMSO has been shown to be an effective solvent for highly hydrophobic compounds.79 Therefore, the polymer was dissolved in THF:MeOH:DMSO at a ratio of 85:10:5 (v:v:v) at a concentration of 0.1 mg/mL. The salt (AgTFA, 10 mg/mL) was added to the solution at a concentration of 0.1%.

DMSO is a supercharging agent. Supercharging agents are typically used to increase the charge state on proteins, in order to improve the analysis by shifting m/z values into a region of higher sensitivity and mass resolution.84 DMSO increases the surface tension of the ESI droplets, allowing the droplet to hold an increased number of charges,104 or in this case increasing the number of charges on the droplets from zero to one.

Ubiquitin (as a standard for IM-MS) was dissolved in 5:5 H2O:ACN at a concentration of 0.68 mg/mL. Twenty uL of acetic acid was added to 1000 uL of the ubiquitin solution

5.2.4 Synapt Q/ToF Parameters

The trap CE for MS/MS analysis is listed with each respective figure. The transfer cell parameters were as follows: wave height – 7.0V, wave velocity – 200 m/s, and gas flow rate – 22.7 mL/min.

69 5.3 Results

The MALDI mass spectrum of the polymer can be seen in Figure 5.1. It confirms the expected repeat unit (theoretical value = 480.0408). An expanded trace (Figure 5.2) indicates the presence of two distributions with two different end groups: 0 Da and 20

Da. The 20 Da end group matches the expected polymer structure (Scheme 5.1), which contains linear branches and a hydroxyl group at the focal point. On the other hand, an end group of 0 is characteristic of a cyclic structure, resulting by the loss of HF. This structure could be a large macrocycle, formed from a completely linear (un-branched) architecture by HF loss, or a tadpole, formed from a branched (or hyperbranched) architecture by HF loss. As can be seen in Figure 5.2, the cyclic distribution is higher in intensity than the linear distribution, which is interesting because the synthesis was intended to make a linear or branched polymer with all-linear chains (as in Scheme 5.1).

The intensities of the cyclic oligomers are lower in the ESI mass spectrum (Figure

5.3), indicating that HF loss and cyclization may be partly caused by MALDI, as the polymer probably absorbs at the laser wavelength for MALDI (355 nm). ESI is a softer ionization method than MALDI. The presence of cyclic species in the ESI spectrum provides evidence that cyclization also occurs during synthesis.

70 Figure 5.1. MALDI mass spectrum of the fluorinated polymer.

All peaks correspond to [M+Ag]+ ions.

71 Figure 5.2. Zoomed-in view of MALDI spectrum showing the two distributions.

All peaks correspond to [M+Ag]+ ions.

The fragmentation pathways of the fluorinated polymer were examined by

MS/MS on the Synapt Q/ToF mass spectrometer. The MS/MS spectrum of the linear 2- mer is shown in Figure 5.4. Plausible structures of the observed fragments can be seen in

Schemes 5.2. The silver ion, a proton, or a carbocation on the molecule provides the charge on the fragments. The proposed structures explain how the m/z values of the

MS/MS fragments arise by simple C-F, C-H, and/or C-O bond cleavages in the [M+Ag]+ precursor ion. Isomerization to more stable structure containing less unpaired electrons is possible, but cannot be ascertained without isotope labeling or high-level electronic structure calculations, which are beyond the scope of this study.

72 designates cyclic n text) designates oligomers still n L Scheme 5.1). C tadpoles, see shown in (structure point oligomers (macrocycles or focal spectrum of the fluorinated polymer. the OH Figure 5.3. ESI Figure containing

73 Figure 5.4. MS/MS spectrum of the [M+Ag]+ ion from the 2-mer of the linear

fluorinated polymer, acquired at a CE of 40 eV.

74 Scheme 5.2. Plausible structures of the MS/MS fragments from the linear 2-mer.

Isomeric structures are possible. Continued on page 76.

75 Scheme 5.2. Continued from page 75.

There are two theoretical structures possible for the 3-mer in the cyclic distribution: a macrocycle and a tadpole (Scheme 5.3). The IM-MS spectrum shows a monomodal drift time distribution (Figure 5.5), hence, only one of these structures is present in the silverated 3-mer analyzed. The IM-MS characteristics of other oligomers from the cyclic distribution are also consistent with the formation of one monomer only.

In order to determine which is formed, MS/MS was utilized. If the cyclic distribution has a tadpole structure, the MS/MS spectrum should show fragments similar to those observed in the linear MS/MS spectrum of the linear 3-mer, because a tadpole also contains a linear tail. Meanwhile, a macrocyclic (circular) isomer would require at least two bond cleavages to form fragments with less repeat units. Such fragmentation must be associated with a higher activation energy than the single bond cleavage needed to

76 break up a linear architecture; hence, the MS/MS spectrum of a macrocycle should be substantially different from that of an isomeric tadpole.62

Figure 5.5. IM-MS spectra for (a) cyclic 3-mer and (b) linear 3-mer

Scheme 5.3. Two possible structures for the cyclic 3-mer.

Figures 5.6 and 5.7 depict the MS/MS spectra of the cyclic and linear 3-mer, respectively. Both spectra were collected at a collision energy of 55 eV, to deposit the same internal energy onto both [M+Ag]+ ions. The MS/MS spectrum of the linear 3-mer shows several major fragment peaks as well as many minor fragment distributions, whereas the MS/MS spectrum of the cyclic 3-mer shows only one major fragment peak and several minor fragment distributions. Based on these differences, the cyclic distribution is more likely to be in the form of the macrocyclic (circular), rather than the tadpole structure. The 178 Da loss is readily accounted for by benzylic C-O bond

78 cleavage within the macrocycle followed by radical-directed elimination of tetrafluoro methylene cyclohexadianone. This fragmentation channel also dominates for the 4-mer

(Figure 5.8), further corroborating the macrocyclic (circular) architecture.

Figure 5.6. MS/MS spectrum of the silverated 3-mer from the cyclic distribution of the

fluorinated polymer, acquired at a collision energy of 55 eV.

79 Figure 5.7. MS/MS spectrum of the silverated 3-mer from the linear distribution of the

fluorinated polymer. (CE = 55 eV)

80 the of distribution from the cyclic silverated 4-mer fluorinated polymer (CE 60 eV). = spectrum of the Figure 5.8. MS/MS Figure

81 Determining the structure of the linear distribution is more complex. There is one structure possible for the 2-mer, two possible structures for the 3-mer (not considering positional isomers), three possible structures for the 4-mer, etc. The MS/MS spectra of the isomeric structures might be similar. Definitive structural assignment is difficult, because reference spectra of pure isomers are not available. Nevertheless, hints about the most likely structure formed can be obtained by careful interpretation of the fragments observed in the MS/MS spectra of the 3-mer (Figure 5.6) and 4-mer (Figure 5.8). The linear and branched architectures possible for the 3-mer are depicted in Scheme 5.4. A

500 Da ion, which gives rise to the base peak in Figure 5.6, can take place in either structure. In contrast, 480 Da and 480 Da losses can only occur from the linear isomer.

Thus, the 3-mer must contain the linear isomer. Whether the branched isomer is coproduced is less clear. A 1000 Da loss is feasible if both branches are cleared. (cf.

Scheme 5.4); however, the same fragment can also be formed by consecutive HF loss after 980 Da elimination (such 20 Da loses are common with fluorinated polymers).

Fortunately, the 4-mer reveals additional insight. It mainly loses a 960 Da moiety (Figure

5.8), which requires that at least 3 repeat units are connected in a linear fashion, cf.

Scheme 5.5. The 480 Da loss again affirms that the product contains 4-mers with all repeat units connected linearly. Small amounts of a branched isomer can, however, not be excluded.

82 Scheme 5.4. Structurally diagnostic MS/MS fragments expected from the linear and branched architecture of the 3-mer. Note that the linear structure has positional isomers, formed by chain propagation at the pentafluorophenyl rings attached at C3 of the central

phenyl group. All bond clevages shown (except the 181 Da loss) are accompanied by

H-rearrangement to the moiety being lost.

83 with linear 4-mer fragments from the architecture. Scheme 5.5. Structurally diagnostic MS/MS Scheme 5.5. Structurally

84 The IM-MS spectrum of the uncyclized 3-mer shows a monomodal drift time distribution, consistent with the formation of one isomer (Figure 5.4b). It is possible, however, that both architectures (linear and branched) are formed, but do not differ sufficiently in their collision cross section (CCS) to be resolved upon IM-MS. This can be found by simulations. Theoretical structures modeled using the Materials Studio software can be used to calculate CCSs with the MOBCAL program. However,

MOBCAL is not parameterized for fluorine atoms. Therefore, theoretical CCSs of the possible structures cannot be determined. In addition, the structure of the linear compound can be deduced from that of the cyclic product (vide infra). Table 5.1 summarizes the experimental CCSs for the oligomers examined.

Table 5.1. Experimental CCS for cyclic and

linear fluoropolymer oligomers.

[M+Ag] m/z DT (a) DT' (b) Ω' (c) Ω (d) Linear (ms) (ms) (Å2) (Å2) 2-mer 1086.99 3.76 3.71 1125 216 3-mer 1567.03 6.75 6.69 1496 285 4-mer 2047.07 9.93 9.87 1806 344 Cyclic 2-mer 1066.99 3.41 3.36 1072 206 3-mer 1547.03 6.42 6.36 1460 279 4-mer 2027.07 9.44 9.38 1762 335

(a) Measured drift time.

(b) Corrected drift time (Equation 2.3).

(d) Experimental CCS (Equation 2.7).

85 y = 595.53x0.4846 Ubiquitin R² = 1 2000 1800 1600 ) 2 1400 Å 1200 1000 800 600 Norm. Norm. CCS( 400 200 0 0 2 4 6 8 10 12 Corr. DT' (ms)

Figure 5.9. Calibration curve for ubiquitin.

The cyclic structure is most likely formed during the synthesis when a linear oligomer wraps around and cyclizes. If the linear structure experiences hyperbranching, a cyclic distribution with tadpole structure would be formed. Since only cyclic distribution with macrocyclic (circular) structure was detected (vide supra), and since such structure can only be generated by HF loss from an all-linear distribution, it is reasonable to assume that the linear distribution contains no branches.

5.4 Conclusions

Using the supercharging agent DMSO enabled ESI of the fluoropolymer and

MS/MS analysis of its distributions to determine their architectures. Based on the

MS/MS spectra, the cyclic distribution was found to consist of a macrocyclic, not tadpole oligomers. Based on that information, along with IM-MS and MS/MS data, the linear oligomers were determined to be unbranched. This means that the monomer used does

86 not form hyperbranched structures at low molecular weight. The bulkiness of the monomer could cause this result.

87 CHAPTER VI

SUGAR-BASED STRUCTURAL ISOMERS

6.1 Introduction

Carbohydrate polymers, also known as glycopolymers, have many potential applications.105 Glycopolymers have been investigated for sensing and targeting lectin,105,106 tethered to gold nanoparticles105 or coated on iron oxide nanoparticles.106

Glycopolymers have also been used as antiadhesives in the prevention of irritable bowel disease,107 for immune modulation,108 and as brushes for specific lectin binding.109 The many practical applications for glycopolymers have created a need for synthetic polymers with sugar pendants.106 The ability to synthesize glycopolymers with a specific sugar isomer would allow for additional specificity. Motivated by this potential, Cesar Lopez in the research group of Dr. Coleen Pugh at the University of Akron synthesized sugar- based monomers with different side-chain functionalities. Several such monomers and non-polymerizable analogs have been characterized by ion mobility mass spectrometry

(IM-MS).

The sugars included in the examined monomers or analogs were α-D- mannopyranose, β-D-mannopyranose, and β-D-glucopyranose (Scheme 6.1). Seven different substituents were attached onto each of the three sugars, as seen in Scheme 6.2, with the exception of R4, with which only the α-D-mannopyranose and β-D- mannopyranose were derivatized.

88 Scheme 6.1. Monosaccharides examined for incorporation into glycopolymers.

Scheme 6.2. Monosaccharide derivatives studied.

Disaccharide and trisaccharide anomers have been previously studied by IM-MS.110 IM-

MS studies on anomeric monosaccharides have only been reported for their methylated derivatives.66,111 This dissertation describes the first such study on monosaccharide anomers with larger substituents. Two of the sugars in Scheme 6.1 are anomers

(epimers), whereas the third one is a diastereomer of the anomeric pair. All are isomers with exactly the same mass.

89 6.2 Experimental Methods

The following sections discuss the sample preparation methods and experimental parameters used in the rest of the chapter.

6.2.1 Synthesis of monomers

The monomers were synthesized by Cesar Lopez in Dr. Pugh’s group at the

Department of Polymer Science, The University of Akron.

6.2.2 ESI Preparation

The samples were prepared for ESI analysis by dissolving them at a concentration of 0.1 mg/mL in 9:1 (v/v) THF: MeOH. One uL of NaTFA solution (at a concentration of 10 mg/mL in THF) was added to the final solution.

6.2.3 Synapt Q/ToF Parameters

The IM-MS cell parameters were as follows: wave height – 7.0V, wave velocity –

200 m/s, and N2 gas flow rate – 22.7 mL/min. The Trap CE was set to 22 eV to induce fragmentation. The wave height and wave velocity on the transfer cell were 7.0 V and

200 m/s, respectively.

6.3 Results

The mass spectra of the monomers are depicted below (Figurse 6.1 – 6.7). The monosaccharide in the samples from which these spectra were acquired was the α-D-

90 mannopyranose isomer. The calculated (theoretical) values for each structure are included in the figure legend, as well as the mass accuracy for each spectrum. All ions observed are [M+Na]+.

Figure 6.1. Mass spectrum of R1-substitued α-mannose.

Calculated mass = 413.1054 Da. Mass accuracy = 123 ppm.

91 Figure 6.2. Mass spectrum of R2-substituted α-mannose. The minor peak at m/z 449

arises from an impurity; the measured m/z corresponds to a sodiated mannose

disaccharide with two acetyl groups (C16H26O13). Calculated mass = 447.0665 Da.

Mass accuracy = 127 ppm.

92 Figure 6.3. Mass spectrum of R3-substituted α-mannose.

Calculated mass = 427.1211 Da. Mass accuracy = 217 ppm.

Figure 6.4. Mass spectrum of R4-substituted α-mannose.

Calculated mass = 425.1054 Da. Mass accuracy = 226 ppm.

93 Figure 6.5. Mass spectrum of α-mannose of the R5 substituted sugar.

Calculated mass = 475.1211 Da. Mass accuracy = 153 ppm.

Figure 6.6. Mass spectrum of α-mannose of the R6 substituted sugar.

Calculated mass = 491.1524 Da. Mass accuracy = 219 ppm.

94 Figure 6.7. Mass spectrum of α-mannose of the R7 substituted sugar.

Calculated mass = 885.2999 Da. Mass accuracy = 181 ppm.

The isotope pattern in each spectrum matches the expected pattern for each monosaccharide, except for the chlorinated sample, which contains a disaccharide impurity (cf. Figure 6.2). The spectra for the β-D-mannopyranose and β-D- glucopyranose isomers are indistinguishable within experimental error; therefore, they have been omitted to avoid redundancy.

In order to distinguish the α-D-mannopyranose, β-D-mannopyranose, and β-D- glucopyranose isomers, MS/MS spectra were acquired. The MS/MS spectra of the R3- substituted monosaccharides can be seen in Figure 6.8.

95 Figure 6.8. MS/MS spectra of the R3-substituted monosaccharides. CE = 22 eV.

Plausible fragmentation pathways leading to the fragments observed are presented in

Scheme 6.3. The first of the major fragments (i.e. m/z 367) is formed by a loss of 60 Da, which corresponds to the loss of an acetyl group in the form of acetic acid via the 1,5-H rearrangement. This reaction produces a double bond inside the saccharide ring. After a consecutive loss of 60 Da via the same mechanism (to form 307 m/z), no more double bonds can be accommodated in the pyranole ring. Thus, the next fragment (m/z) involves the loss of 42 Da, which converts an acetyl group still attached to the ring to a hydroxyl group. An additional sequence of fragmentations occurs after all acetyl pendants have been eliminated. These are rationalized in the lower section in Scheme 6.3. The MS/MS

96 spectra of the monosaccharide isomers are identical; consequently fragmentation cannot be used to distinguish the isomers.

Scheme 6.3. Fragmentation pathways accounting for the MS/MS fragments from the R3-

substituted sugar monosaccharide.

IM-MS was then tested as a means to differentiate the isomers. The ion mobility spectra for each of the monosaccharides can be seen in Figures 6.9 – 6.15.

97 Figure 6.9. IM-MS spectra of the R1-substituted monosaccharides: (a) α-D-