NATURAL RUBBER BIOSYNTHESIS: PERSPECTIVES from POLYMER CHEMISTRY a Dissertation Presented to the Graduate Faculty of the Univers

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NATURAL RUBBER BIOSYNTHESIS:

PERSPECTIVES FROM POLYMER CHEMISTRY

A Dissertation

Presented to

The Graduate Faculty of The University of Akron

In Partial Fulfillment

of the Requirements for the Degree

Doctor of Philosophy

Cheng Ching Kurt Chiang

December, 2013 NATURAL RUBBER BIOSYNTHESIS:

PERSPECTIVES FROM POLYMER CHEMISTRY

Cheng Ching Kurt Chiang

Dissertation

Approved: Accepted:

Advisor Department Chair Dr. Judit E. Puskas Dr. Coleen Pugh

Committee Member Dean of the College Dr. Abraham Joy Dr. Stephen Z. D. Cheng

Committee Member Dean of the Graduate School Dr. Matthew Becker Dr. George R. Newkome

Committee Member Date Dr. Chrys Wesdemiotis

Committee Member Dr. Peter Rinaldi

ABSTRACT

Natural Rubber (NR) is an important strategic raw material for manufacturing a

wide variety of industrial products. NR has been mainly obtained from Hevea

brasiliensis. The USA is self-sufficient in the production of synthetic rubber while NR supply in USA is mainly imported from Southeast Asia. However, synthetic rubber cannot match the performance of imported Hevea NR. It is a matter of great concern that the USA is highly vulnerable to disruptions of NR supply because of a possible introduction of leaf blight into plantations as none of the trees in plantations across

Southeast Asia have resistance to blight.

Puskas et al. postulated that the biosynthesis of polyisoprenoids in general, and that of NR in particular, may proceed by a living carbocationic polymerization process.

“Natural living carbocationic polymerization” (NLCP) mechanism was proposed in terms of accepted polymer chemical formalism, i.e., initiation, propagation, and equilibria between active and dormant species. This thesis studied the fundamental steps of NR biosynthesis in two ways: 1) characterization of NR from stabilized latex and in vitro

biosynthesis; 2) addition of isoprene (IP) and other derivatives to shift the enzymatic

equilibrium of the terpenoid biosynthesis lifecycle. Synthetic initiators to be used in in

vitro NR biosynthesis system were prepared and characterized.

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For the first portion of this thesis, in vitro NR biosynthesis was studied using high

resolution size exclusion chromatography (HR-SEC). Then, various compounds such as

isoprene (IP) and amylene were introduced to the in vitro NR biosynthesis system. The rationale behind these studies was based on the terpenoid biosynthesis lifecycle. HR-

SEC, gravimetric analysis, in situ Raman spectroscopy and NMR spectroscopy were used to study scenarios where the equilibrium was disrupted by the introduction of foreign chemical compounds.

The second goal of this research was to design and synthesize a synthetic initiator for the future synthesis in vitro of a novel polyisobutylene-block-cis-1,4-polyisoprene

(PIB-b-NR) diblock copolymer. Two feasible synthetic pathways were developed to yield the target synthetic initiator: one by conventional syntheses and the other through an enzymatic pathway. The ultimate purpose of this synthetic initiator is to explore if the cis-prenyltransferase enzyme will recognize the compound and eventually apply this concept to novel terpenoids/NR in vitro. A seven-step synthetic strategy for the preparation of the PIB-NPP synthetic initiator is proposed and presented.

DEDICATION

To my parents, Wei Ching and Li Chen Chiang for all their love and support.

v ACKNOWLEDGEMENTS

I would like to express my gratitude to my advisor Professor Judit E. Puskas for

her guidance and support throughout the course of this research. I would also like to

thank my graduate committee members Dr. Matthew L. Becker, Dr. Abraham Joy, Dr.

Chrys Wesdemiotis and Dr. Peter Rinaldi for their helpful comments and objective

criticisms. Also, I would like to thank Dr. Balaka Barkakaty for her extended amount of

her help and time.

I am grateful to Dr. Puskas’ team members for their collaboration and for providing useful feedback throughout this research. I would like to especially thank Dr.

Mustafa Yasin Sȩn, Dr. Elizabeth Foreman-Orlowski, Dr. Serap Hayat-Soytaş, Dr. Lucas

Dos Santos, and Dr. Andrey Malkovsky for helpful discussions about my research and laboratory techniques/instrumentations. I would also want to thank Dr. Colleen

McMahan, Dr. Wenshuang Xie, Dr. Alexi Sokolov and Dr. Alain Deffieux for their samples and insights.

I would like to acknowledge financial support by NSF-CHE#0616834 (GOALI) and the Goodyear Tire & Rubber Company.

Finally, it would have been impossible to write this dissertation without my family’s love and support especially from Dr. Ozlem Turkarslan and Dr. Yi-Hsin Weng.

vi TABLE OF CONTENTS

Page

LIST OF TABLES ...... xii

LIST OF FIGURES ...... xiv

CHAPTER

I. INTRODUCTION ...... 1

II. BACKGROUND ...... 7

2.1. Chemical Structure of Natural Rubber ...... 7

2.2. History of Natural Rubber ...... 13

2.2.1. Synthetic Polyisoprenes ...... 15

2.3. Natural Rubber Biosynthesis ...... 19

2.3.1. Rubber Producing Plants ...... 19

2.3.2. Anatomy of the Hevea NR Latex ...... 21

2.3.3. Biochemical Pathway of Natural Rubber Biosynthesis In Vivo ...... 22

2.3.4. Prenyltransferases ...... 28

2.3.5. Mechanism of Prenylation in Short Chain Isoprenoids ...... 29

2.3.6. Proposed Mechanism of Natural Rubber Biosynthesis: Natural Living Carbocationic Polymerization (NLCP) ...... 30

2.4. In Vitro Natural Rubber Biosynthesis ...... 34

2.5. In Vitro Biosynthesis Using Modified Synthetic Initiators ...... 40

vii 2.6. Biomimetic Polymerization of IP ...... 42

III. EXPERIMENTAL ...... 46

3.1. Materials ...... 46

3.1.1. Preparation of Washed Rubber Particles (WRP) ...... 47

3.2. Procedures ...... 48

3.2.1. In Vitro Natural Rubber Biosynthesis (Hevea WRP) ...... 48

3.2.1.1. Small Scale Synthesis (USDA) ...... 48

3.2.1.2. Large Scale Synthesis (USDA) ...... 49

3.2.1.3. “Bioemulative” Experiments Using Synthetic Isoprene with WRP-3 (USDA) ...... 49

3.2.2. Experiments with IAC40 Latex ...... 50

3.2.2.1. Solids Content Determination for Latex and WRP ...... 50

3.2.2.2. In Situ Raman Monitoring ...... 50

3.2.2.3. Micro-Raman Spectroscopy ...... 51

3.2.2.4. Experiments under CO2 Atmosphere ...... 52

3.2.2.5. “Bioemulative” Experiments Using Deuterated Isoprene ...... 53

3.3. Synthesis of Macroinitiator ...... 54

3.3.1. Synthesis of Protected Nerol (PN, product 2) ...... 54

3.3.2. Synthesis of Protected Nerol-OH (PN-OH, product 3) ...... 54

3.3.3. Synthesis of Protected Nerol Tosylate (PN-Ts, product 4) ...... 55

3.3.4. Synthesis of Polyisobutylene-Protected Nerol (PIB-PN, product 5) ...56

3.3.5. Synthesis of Polyisobutylene-Nerol (PIB-Nerol, product 6) ...... 57

3.3.6. Synthesis of Polyisobutylene-Nerol-Bromide (PIB-Nerol-Br, product 7) ...... 57

3.3.7. Synthesis of Tris(tetra-n-butylammonium) Hydrogen Pyrophosphate [(NBu4)3HP2O7] ...... 57

viii 3.3.8. Synthesis of Nerol-PP (Model Reaction for Product 8) ...... 58

3.3.9. Synthesis of Protect Nerol-Divinyl Adipate (PN-DVA, Product 9) ....58

3.3.10. Synthesis of Trimethyl Pentyl Chloride (TMPCl, Product 10) ...... 59

3.3.11. Synthesis of Allyl Trimethyl Pentane (TMP-allyl, Product 11) ...... 59

3.3.12. Synthesis of Trimethyl Pentane-OH (TMP-OH, Product 12) ...... 60

3.3.13. Synthesis of Protected Nerol-Divinyl Adipate-Trimethyl Pentane (PN-DVA-TMP, Product 13) ...... 60

3.3.14. Synthesis of Nerol-Divinyl Adipate-Trimethyl Pentane (Nerol-DVA-TMP, Product 14) ...... 61

3.3.15. Synthesis of Nerol-Bromide-Divinyl Adipate-Trimethyl Pentane (Nerol-Br-DVA-TMP, Product 15) ...... 61

3.3.16. Synthesis of Nerol-Pyrophosphate-Divinyl Adipate-Trimethyl Pentane (Nerol-PP-DVA-TMP, Product 16) ...... 62

3.3.17. Synthesis of Nerol-P(O)(OEt)2 (Model Reaction for Phosphorylation) ...... 62

3.3.18. Synthesis of Nerol-P(O)(OEt)2-DVA-TMP (Product 15) ...... 63

3.3.19. Synthesis of Nerol-PP-DVA-TMP (Product 16) ...... 63

3.4. Laboratory Techniques and Instrumentation ...... 64

3.4.1. Air-free Technique ...... 64

3.4.2. Thin Layer Chromatography (TLC) ...... 64

3.4.3. Column Chromatography ...... 65

3.4.4. Size Exclusion Chromatography (SEC) ...... 66

3.4.5. NMR Sample Preparation ...... 67

3.4.6. 1H NMR Procedure ...... 68

3.4.7. 13C NMR Procedure ...... 68

3.4.8. Gas Chromatography (GC) ...... 68

ix 3.4.9. Matrix Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry (MALDI-ToF MS) ...... 69

3.4.10. Electrospray Ionization Mass Spectrometry (ESI-MS) ...... 69

IV. RESULTS AND DISCUSSION ...... 71

4.1. In Vitro Natural Rubber Biosynthesis ...... 71

4.1.1. Monitoring the Growth of in vitro Natural Rubber by High- Resolution Size Exclusion Chromatography (HR-SEC) ...... 71

4.2. Substitution of the Isopentenyl Pyrophosphate (IPP) Monomer with Synthetic Isoprene (IP) ...... 82

4.2.1. Monitoring the Effects of IP in in vitro Natural Rubber Biosynthesis. .83

4.2.2. Incubation of Synthetic D-IP with Hevea Latex...... 102

4.2.3. In Vitro NR Biosynthesis under CO2 Atmosphere...... 108

4.2.4. In Vitro NR Biosynthesis in the Presence of Amylene ...... 111

4.3. In Situ Micro-Raman Monitoring of In Vitro Natural Rubber (NR) Biosynthesis ...... 113

4.3.1. Raman Monitoring of In Vitro NR Biosynthesis Between Glass Slides ...... 113

4.3.2. Raman Monitoring of In Vitro NR Biosynthesis with Micro- Cavity Slides ...... 115

4.3.3. Raman Monitoring in Sealed Silanized Vials ...... 116

4.3.4. Raman Monitoring with Deuterated-Isoprene (D-IP) ...... 124

4.3.5. Raman Monitoring CO2 Atmosphere ...... 124

4.3.6. Effect of Addition on Latex Particle Size ...... 126

4.4. Macroinitiator Synthesis ...... 128

4.4.1. Synthesis of Protected Nerol (PN, Product 2) ...... 131

4.4.2. Synthesis of Protected Nerol-OH (PN-OH, Product 3) ...... 134

4.4.3. Synthesis of Protected Nerol Tosylate (PN-Ts, Product 4) ...... 136

x 4.4.4. Synthesis of Polyisobutylene Protected Nerol (PIB-PN, Product 5) .138

4.4.5. Deprotection of PIB-PN to PIB-Nerol (Product 6) ...... 142

4.4.6. Synthesis of Polyisobutylene-Nerol-Bromide (PIB-Nerol-Br, Product 7) ...... 144

4.4.7. Synthesis of Nerol Pyrophosphate (Nerol-PP, Model Reaction for Product 8) ...... 146

4.4.8. Synthetic Stretegy to Yield Macroinitiator Using Enzyme Catalysis ...... 147

4.4.9. Synthesis of Protected Nerol-Divinyl Adipate (PN-DVA, Product 9) ...... 149

4.4.10. Synthesis of TMP-OH (Product 12, PIB dimer) ...... 153

4.4.11. Synthesis of PN-DVA-TMP (Product 13) ...... 159

4.4.12. Synthesis of Nerol-DVA-TMP (Product 14) ...... 162

4.4.13. Synthesis of Nerol-Br-DVA-TMP (Product 15) ...... 165

4.4.14. Synthesis of Nerol-PP-DVA-TMP (Product 13) (Chen’s Method) ..167

4.4.15. Synthetic scheme to yield Nerol-OPP-DVA-TMP (Product 16) (Coates’Method) ...... 169

4.4.16. Synthesis of Nerol-OP(O)(OEt)2 (Model Reaction) ...... 170

4.4.17. Synthesis of Nerol-PP (Model Reaction) ...... 174

4.4.18. Synthesis of Nerol-OP(O)(OEt)2-DVA-TMP (Product 15) ...... 176

4.4.19. Synthesis of Nerol-PP-DVA-TMP (Product 16) ...... 179

V. CONCLUSIONS ...... 185

REFERENCES ...... 188

APPENDIX ...... 201

xi LIST OF TABLES

Table Page

2.2.1 Microstructures in synthetic PIPs ...... 15

4.1.1 HR-SEC data of WRP-1...... 75

4.1.2 Approximate MW of peaks 1L to 5L from HR-SEC...... 78

4.1.3 Gravimetric analysis of WRP-3/24...... 80

4.1.4 Gel fraction analysis of WRP-3 and WRP-3/24 ...... 80

4.1.5 SEC data of WRP-3 and WRP-3/24 ...... 81

4.2.1 Experimental conditions for WRP-3 with IP ...... 83

4.2.2 Gravimetric analysis summary for experiments with WRP-3 ...... 83

4.2.3a IAC40 latex solid content determination by freeze drying ...... 86

4.2.3b IAC40 WRP solid content determination by freeze drying ...... 86

4.2.4a Solid content of IAC40 latex after washing the coagulated rubber ...... 86

4.2.4b Solid content of IAC40 WRP after washing the coagulated rubber ...... 86

4.2.5a Solid content of IAC40 latex obtained by precipitation in methanol ...... 87

4.2.5a Solid content of IAC40 WRP obtained by precipitation in methanol ...... 87

4.2.6 Summary gravimetric data of in vitro NR samples ...... 89

4.2.7 Gel content of in vitro NR samples ...... 90

4.2.8 High and low MW parts of the soluble fractions obtained from the IAC40 latex and WRP ...... 92

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4.2.9a SEC analysis of the soluble fractions obtained from IAC40 latex before and after incubation with IP ...... 94

4.2.9b SEC analysis of the soluble fractions obtained from IAC40 WRP before and after incubation with IP ...... 94

4.2.10 Gravimetric summary of D-IP experiments ...... 106

4.2.11 Gravimetric analysis of in vitro NR biosynthesis under CO2 atmosphere ...... 109

4.2.12 Gel content of in vitro NR samples ...... 109

4.2.13 SEC analysis of the soluble fractions obtained from IAC40 latex/WRP before and after incubation with IP in CO2 ...... 111

4.2.14 Gravimetric summary for Amylene experiments ...... 112

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LIST OF FIGURES

Figure Page

2.1.1 DL-limonene (a) and isoprene (b) ...... 7

2.1.2 Suggested building blocks of NR before (a) and after (b) 1910 ...... 8

2.1.3 Chemical structure of NR. Head (ω) and end group (α) structure is not proven ...... 9

2.1.4 13C NMR spectrum of NR from Hevea (a) and Guayule (b) ...... 10

2.1.5 13C NMR spectrum of NR from L. volemus ...... 11

2.1.6 Deproteinization by treatment with 1~2% ethanol. (DMAPP denotes dimethylallyl pyrophosphate, FPP denotes Farnesyl pyrophosphate.) ...... 12

2.2.1 PIP microstructures. *the cyclics shows an example of C15, other cyclics may occur ...... 16

2.2.2 NMP of IP using 2,2,5-trimethyl-4-phenyl-3-azahexane-3oxy-nitroxide as initiator ...... 17

2.3.1 Examples of rubber producing plants ...... 20

2.3.2 Visualization of NR particles and its structure ...... 21

2.3.3 Structure of isopentenyl pyrophosphate IPP at pH = 7.4 ...... 22

2.3.4 Terpenoids biosynthesis cycle ...... 24

2.3.5 Synthesis of NR in H. brasiliensis. OPP stands for the pyrophosphate end-group and HOPP represents pyrophosphoric acid ...... 25

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2.3.6 Structure of the allylic oligoisoprene pyrophosphates. DMAPP: dimethyl allyl pyrophosphate, GPP: geranyl pyrophosphate, FPP: farnesyl pyrophosphate, GGPP: geranylgeranyl pyrophosphate ...... 26

2.3.7 Biochemical representation of rubber biosynthesis ...... 27

2.3.8 A scheme of the active sites in avian trans-prenyltransferase. The oval represents a large motive proposed to stop chain growth ...... 28

2.3.9 The mechanism of terpenoid biosynthesis proven by Poulter et al ...... 30

2.3.10 Proposed NLCP mechanism of NR biosynthesis ...... 31

2.3.11 Yokozawa’s concept of chain-growth polycondensation ...... 32

2.3.12 Attempted “bio-inspired” synthesis of cis-1,4-polyisoprene. (The resulting polymer was in the trans conformation) ...... 33

2.4.1 Lineweaver-Burk Plot of IPP concentration vs. reaction velocity (V) in the determination of the enzymatic activity of Guayule WRP ...... 35

2.4.2 SEC trace of Hevea BF (dashed line) and in vitro NR (solid line) ...... 36

2.4.3 in vitro guayule NR biosynthesis ...... 38

2.4.4 Experimental MW results from Cornish et al.’s in vitro NR system for three types of WRPs (Fig tree, Guayule, and Hevea) a) 0.25 µM FPP, b) 2.5 µM FPP ...... 39

2.5.1 Chemical structure of DATFP-GPP ...... 41

2.5.2 Chemical structure of di-isobutylene-neryl pyrophosphate ...... 42

2.6.1 Biomimetic initiation of IP polymerization from Puskas et al ...... 43

2.6.2 Possible carbocationic polymerization pathways for IPOH ...... 44

2.6.3 Chemical scheme of cationic polymerization of IP with 1-(4-methoxyphenyl) ethanol as the initiator and B(C6F5)3 as the co-initiator .....45

3.2.1 Experimental set-up for the in situ Raman measurements ...... 51

3.2.2 Micro-Raman instrumentation in the Sokolov lab: a) before the experiment b) during in situ monitoring ...... 52

3.2.3 Methodology to exchange the atmosphere within the closed vial ...... 53

3.4.1 High resolution SEC system at Puskas Lab ...... 67

4.1.1 Micro-well plates in which in vitro NR biosyntheses are performed ...... 72

4.1.2 An example micro-well and its constituents a) during and b) after incubation ...... 73

4.1.3 Enzyme activity measurement of RRIM 600 WRP using 14C IPP (USDA) ...... 73

4.1.4 SEC RI trace of WRP-1 ...... 74

4.1.5 Conformation Plot of WRP-3 ...... 75

4.1.6 SEC of WRP-1, WRP-1/5 and WRP-1/24. High MW region a) LS trace, b) RI trace ...... 77

4.1.7 SEC RI trace of WRP-1 and WRP-1/24. Low MW region ...... 77

4.1.8 MALDI-ToF spectrum of fractionated low MW Hevea NR (H600 clone) ...... 79

4.1.9 SEC traces of WRP-3 before and after incubation (a) RI traces of low MW region, (b) RI traces of high MW region ...... 81

4.2.1 Enzyme activity measurement using 14C-labelled IPP (USDA) ...... 85

4.2.2 Zoomed SEC RI chromatograms of the soluble fractions from IAC40 and WRP for comparison: a) high MW region, b) low MW region ...... 91

4.2.3 SEC RI trace of KC_092309_L_IP1 and starting latex ...... 93

4.2.4 Difference between (a) least square fit (unmodified) and (b) least absolute residual fit (modified) ...... 95

4.2.5 1H NMR (500 MHz) spectrum of KC_092309_L_IP1: 0-3.5 ppm region (Concentration: 10 mg/mL, 128 scans, d1 = 10 sec, Pulse angle = 90o, o T = 25 C, Solvent: toluene-D8.) ...... 97

4.2.6 1H NMR (500 MHz) spectrum of KC_092309_L_IP1: 3.75-7.5 ppm region (Concentration: 10 mg/mL, 128 scans, d1 = 10 sec, o o Pulse angle = 45 , T = 25 C, Solvent: toluene-D8.) ...... 97

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4.2.7 13C NMR spectrum of KC_092309_L_IP1: 0-50 ppm region. (Concentration: 20 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: Chloroform-D.) ...... 98

4.2.8 13C NMR spectrum of KC_092309_L_IP1: 100-150 ppm region (Concentration: 20 mg/mL, 10,000 scans, d1 = 10s, Pulse Angle = 90o, T = 25°C, Solvent: Chloroform-D.) ...... 99

4.2.9 13C NMR (125 MHz) spectrum of KC_100909_L_IP5: 0-50 ppm region (Concentration: 50 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6.) ...... 100

4.2.10 13C NMR (125 MHz) spectrum of KC_100909_L_IP5: 100-155 ppm region (Concentration: 50 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6.) ...... 100

4.2.11 13C NMR (125 MHz) spectrum of KC_102109_W_IP8: 0-50 ppm region (500 MHz Concentration: 60 mg/mL, 10,000 scans, d1 = 10s, o Pulse angle = 90 , T = 25°C, Solvent: benzene-D6.) ...... 101

4.2.12 13C NMR (125 MHz) spectrum of KC_102109_W_IP8: 95-155 ppm region (Concentration: 60 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6.) ...... 102

4.2.13 Comparison of 1H NMR spectra of IP (red) and D-IP (black) (300 MHz, Concentration: 20 mg/mL, 32 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 103

4.2.14 Comparison of the 13C NMR spectra of IP (red) and D-IP (black) (300 MHz, concentration: 20 mg/mL, 32 scans, d1 = 10s, o Pulse angle = 90 , T = 25°C, Solvent: benzene-D6.) ...... 104

4.2.15 GC chromatograms of IP (red) and D-IP (black) ...... 105

4.2.16 13C NMR spectra (500 MHz) of the D-IP experimental series, 091610_D-IP_latex 2 (top), 092910_L_IP1_50D-IP_50IP (middle) and 092910_L_100IP (bottom), 0-140 ppm region. 10,000 scans, o d1 = 10s, Pulse angle = 90 , Solvent: benzene-D6, (Concentration: 15 mg/mL for KC091610_L_IP1(100D-IP) and 50 mg/mL for both KC092910 experiments.) ...... 107

4.2.17 SEC comparison RI traces of KC_102709_Latex_IAC40 and KC_121109_L_IP1(CO2/50X)...... 110

xvii

4.2.18 SEC comparison RI traces of KC120909_IAC40_WRP and KC121409_W_IP3 (CO2/50X)...... 110

4.2.19 Chemical structure of amylene...... 111

4.2.20 SEC comparison RI traces of 102709_Latex and KC_101510_L_Amy1 observed after incubation of IAC40 NR latex with amylene. The other two experiments (KC101510_L_Amy2 and KC101510_L_Amy3, not shown) showed similar results with KC101510_L_Amy1 with no changes in the SEC traces...... 112

4.3.1 Signals attributed to the C=C Raman-active vibrations in PIP and IP (Isoprene: liquid IP with stabilizer; Mixed sample: WRP RRIM600+IP 2/1 (w/w))...... 114

4.3.2 The schematics of the bottle illuminated by the Raman beam...... 117

4.3.3 Fluorescence problems illustrated for KC_092309_L_IP1 with 80X objective (092309_L_WRP: 0.5340g IAC40 latex, 0.1022g IP, 24h, RT) ...... 118

4.3.4 Raw spectra for a sequence of data points of KC_102109_W_IP8 (10 min between each spectra, 0.5079g IAC40 WRP, 0.1022g IP, 24h, RT.) ...119

4.3.5 Normalized PIP formation plots for two repeat experiments using 50X. objective (KC_110509_WRP_IP9: 0.5028g WRP, 0.1022g IP, 24h, RT. KC_120209_WRP_IP13: 0.5097g WRP, 0.1021g IP, 24h, RT.) ...... 120

4.3.6 Normalized PIP formation plots for two repeat experiments with 50X objective with error bars. (KC_102309_WRP_IP8: 0.5079g WRP, 0.1022g IP, 24h, RT. KC_111209_WRP_IP10: 0.5843g WRP, 0.1022g IP, 24h, RT.) ...... 121

4.3.7 Normalized IP consumption plots for three repeat experiments with 50X objective (103209_WRP_IP8: 0.5079g WRP, 0.1022g IP, 24h, RT 110509_WRP_IP9: 0.5028g WRP, 0.1022g IP, 24h, RT 111209_WRP_IP10: 0.5843g WRP, 0.1022g IP, 24h, RT.) ...... 122

4.3.8 Raman monitoring of in vitro NR biosynthesis. (KC_102109_WRP_IP8: 0.5079 g IAC40 WRP, 0.1021 g IP, 24 hrs, RT) ...... 123

4.3.9 Raman monitoring of KC091710_L_IP3 (100DIP) ...... 124

4.3.10 PIP growth comparison between WRP experiments and KC_121409_W_IP3 (CO2/50X) 0.5085 g IAC40 WRP, 0.102 g IP, o 24 hrs under CO2 atmosphere, at 25 C, color: yellow ...... 125

xviii

4.3.11 PIP growth comparison between KC_092909_L_IP4 and KC_120809_L_IP1 (CO2/50X), 0.6406 g IAC40 Latex, 0.130 g IP, 24 hrs under CO2 atmosphere at 25 C, color: yellow ...... 126

4.3.12 Optical images of Hevea latex particles (Neotex HA) casted on a flat glass slide...... 127

4.3.13 Size distribution of Neotex HA latex particles...... 127

4.4.1 Synthetic strategy to produce PIB-NPP macroinitiator PPTs = pyridinium p-toluenesulfonate; DHP = dihydropyran; NBS = N-bromosuccinimide; TsCl = tosyl chloride; IPA = isopropyl alcohol ...130

4.4.2 1H NMR spectrum of nerol. (300 MHz, 32 scans, d1 = 1s, Pulse angle = 45o , T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.2) ...... 132

4.4.3 1H NMR spectrum of PN (product 2). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.3) ...... 133

4.4.4 13C NMR spectrum of PN (product 2). (500 MHz, 5,000 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.4) ...... 134

4.4.5 1H NMR spectrum of PN-OH (product 3). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.5) ...... 135

4.4.6 13C NMR spectrum of PN-OH (product 3). (300 MHz, 5,000 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.6) ...... 136

4.4.7 1H NMR spectrum of PN-Ts (product 4). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.7)...... 137

4.4.8 13C NMR spectrum of PN-Ts (product 4) (500 MHz, 5,000 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.8) ...... 138

4.4.9 1H NMR spectrum of starting PIB-OH (#16) (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.9) ...... 139

4.4.10 13C NMR spectrum of starting PIB-OH (#16) (300 MHz, 5,000 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.10) ...... 140

4.4.11 1H NMR spectrum of PIB-PN (product 5) (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.11) ...... 141

xix

4.4.12 13C NMR spectrum of PIB-PN (product 5). (500 MHz, 6,400 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.12) ...... 142

4.4.13 1H NMR spectrum of PIB-Nerol (product 6) (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.13) ...... 143

4.4.14 13C NMR spectrum of PIB-Nerol (product 6) (300 MHz, 5,000 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.14) ...... 144

4.4.15 1H NMR spectrum of PIB-Nerol-Br (product 7) (300 MHz, 128 scans, d1 = 10s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.15) ...... 145

4.4.16 13C NMR spectrum of PIB-Nerol-Br (product 7) (500 MHz, 4,600 scans, d1 = 10s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.16) ...... 146

4.4.17 1H NMR spectrum of Nerol-PP(model compound of product 8) (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D.) ...... 147

4.4.18 Enzyme-catalyzed synthetic scheme to yield Nero-PP-DVA-TMP ...... 149

4.4.19 1H NMR spectrum of PN-DVA (product 9). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D.) ...... 151

4.4.20 13C NMR spectrum of PN-DVA (product 9). (500 MHz, 2,800 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D.) ...... 152

4.4.21 ESI-MS spectrum of PN-DVA (product 9). (solvent: THF, cationizing agent: NaTFA, sample/salt: 100/1 (v/v)) ...... 153

4.4.22 Synthetic scheme to yield TMP-OH (PIB dimer) ...... 154

4.4.23 1H NMR spectrum of TMP-1. (300 MHz, 32 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D.) ...... 155

4.4.24 1H NMR spectrum of TMPCl. (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D.) ...... 156

4.4.25 1H NMR spectrum of TMP-allyl. (300 MHz, 32 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D.) ...... 157 4.4.26 1H NMR spectrum of TMP-OH. (300 MHz, 32 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D.) ...... 158

4.4.27 ESI-MS of TMP-OH. (solvent: Methanol, cationizing agent: NaTFA, sample/salt: 100/1 (v/v)) ...... 159

4.4.28 1H NMR spectrum of PN-DVA-TMP (product 13). (300 MHz, 32 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.28) ...... 160

4.4.29 13C NMR spectrum of PN-DVA-TMP (product 10). (125 MHz, 3,600 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.29) ...... 161

4.4.30 ESI-MS spectrum of PN-DVA-TMP (product 10). (solvent: THF, cationizing agent: NaTFA, sample/salt: 100/1 (v/v)) ...... 162

4.4.31 1H NMR spectrum of Nerol-DVA-TMP (product 14). (300 MHz, 64 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.31) ...... 164

4.4.32 13C NMR spectrum of Nerol-DVA-TMP (product 14). (75 MHz, 3600 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.32) ...... 165

4.4.33 1H NMR spectrum of Nerol-Br-DVA-TMP (product 15). (300 MHz, 64 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.33) ...... 166

4.4.34 13C NMR spectrum of Nerol-Br-DVA-TMP (product 15). (75 MHz, 3600 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.34) ...... 167

4.4.35 31P NMR spectrum of diethyl chlorophosphate. Bu denotes n-butyl group (300 MHz, 300 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 168

4.4.36 Synthetic scheme to yield Nero-PP-DVA-TMP adapted from Coates et al ...... 170

4.4.37 31P NMR spectrum of diethyl chlorophosphate. (300 MHz, 400 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 171

31 4.4.38 P NMR spectrum of crude product of Nerol-P(O)(OEt)2.(300 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 172

31 4.4.39 P NMR spectrum of Nerol-P(O)(OEt)2. (after column) (300 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 173

xxi

1 4.4.40 H NMR spectrum of Nerol-P(O)(OEt)2. (300 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.40) ...... 174

4.4.41 31P NMR spectrum of Nerol-PP. (crude, Bu denotes n-Butyl) (300 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 175

4.4.42 ESI-MS spectrum of Nerol-PP. (Negative Mode. solvent: acetonitrile, ionizing agent: n/a)………………………………………………………………………176

31 4.4.43 P NMR spectrum of Nerol-OP(O)(OEt)2-DVA-TMP. (300 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 177

1 4.4.44 H NMR spectrum of Nerol-P(O)(OEt)2-DVA-TMP. (300 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.44) ...... 178

4.4.45 31P NMR spectrum of Nerol-PP-DVA-TMP. (product 16) (300 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D.) ...... 180

4.4.46 1H NMR spectrum of Nerol-PP-DVA-TMP. (product 16) (300 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.46) ...... 181

4.4.47 13C NMR spectrum of Nerol-PP-DVA-TMP (product 16). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D. Appendix: B.4.4.47) ...... 182

+ 4.4.48 ESI-MS spectrum of Nerol-PP-TMP-DVA. Bu4N denotes tetra-n-butylammonium ion.(Solvent: acetonitrile, cationizing agent: n/a) ...... 184

xxii

CHAPTER I

INTRODUCTION

Plants are present in all aspects of everyday life; they provide food, produce the oxygen that organisms breathe, and serve as raw materials in many of our clothes, buildings, drugs and perfumes. Despite the diversity of plants, human life relies only on relatively few species. Even though the Olmec, who are often referred to as the “rubber people”, first discovered Natural Rubber (NR) around 1,600 B.C.1, it was not until 1839 that rubber had its first practical application in the industry when Charles Goodyear accidentally dropped rubber and sulfur on a hot stovetop, causing it to be resilient like leather while retaining elasticity.2, 3 NR is a critical and strategic industrial raw material

for manufacturing a wide variety of products, ranging from medical devices and personal

protective equipment to aircraft tires. Car tires are made from 12-14 different rubbers, of

which up to 50% of the rubber used can be NR. Aircraft and race tires contain only NR.2

It is known that more than 2,500 plant species produce NR; however, NR

harvested from a single species, Hevea brasiliensis (Brazilian rubber tree) is the most

important commercial source.4 Today 85% of the USA NR supply is imported from

Southeast Asia and the remaining 15% comes from the South and Central Americas.5, 6 In

2006, the total global production of natural rubber was ~9 million metric tons.7

The US is highly vulnerable to disruptions of NR supply such as a possible

introduction of leaf blight into plantations.2 None of the trees in plantations across

Southeast Asia have resistance to blight. South American leaf blight (SALB) is a fungal disease common to rubber trees caused by the endemic fungus Microcyclus ulei, an

ascomycota (sac fungi) that spreads by spores.7 Currently, SALB is restricted to South

and Central Americas due to strict control over air traffic and freight shipments from

South America to Southeast Asia. However, future intercontinental passenger flights

between Southeast Asia and South America can be detrimental to the rubber plantations

if not controlled properly.

The rapid development of Brazil in the late 19th Century was similar to the Gold

Rush in Alaska. The Industrial Revolution increased the production of automobiles and

spread electricity, which raised the demand of NR significantly and NR became a global

commodity.8 Immigrants flooded to towns such as Belem and Manaus in Northern Brazil

to work in the Amazon Basin. Buildings such as the Amazonas Theater symbolize the

once prosperous Northern Brazil from the “Rubber Boom”. The output of NR from Brazil

was approximately 42,000 tons a year in the late 19th Century, which accounted for

almost the entire global NR market.8 In the early 20th Century, NR production from

British Colonies in Southeast Asia entered the market and devastated the Brazilian NR

economy.9

The success of the Brazilian NR was the envy of the entire world and the British

smuggled H. brasiliensis seeds from Amazon to London in 1876.9 The British scientists developed a higher-yield, more disease resistant variety of H. brasiliensis through grafting and planted them in newly established plantations in the Malaysian Peninsula. 2

The prices of NR from Southeast Asia were significantly cheaper than their Brazilian

counterparts. The main reason for this was because NR produced in Southeast Asia were

planted approximately four meters apart while the NR from Brazil often required the

farmer to walk up to a kilometer between trees.9 The fierce competition between

Brazilian and Southeast Asia NR brought the end of the economic boom of Northern

Brazil.

The Brazilian government did not leave the NR industry without a final attempt to revive Northern Brazil. The Brazilian government partnered with Ford to form Brazil’s

own rubber plantation; over 200,000 trees were planted over 3,200 hectares.7 This

ambitious project was abandoned and posted huge losses with the first reported epidemic

of H. brasiliensis in 1933 by SALB.10 Other attempts to establish a plantation in Brazil or

South America had failed due to the fact that the trees were destroyed before reaching

physiological maturity.

SALB is a rubber tree disease caused by the fungus Microcyclus ulei, an

ascosmycete (sac fungi) that is native to the Amazon area. Modern attempts to control

this disease by chemical (fungicides)11 and biological (breeding and selection)12, 13 methods have failed. As the name implies from SALB, Microcyclus ulei is known to infect young leaves and stems of H. brasiliensis; it was also observed that matured leafs of H. brasiliensis are resistant to M. ulei. In most cases, M. ulei attacks the young leafs of

Hevea, causes grey lesions on the leaf and prevents photosynthesis of these leafs.12 In other words, the development of commercial-scale rubber plantation cannot be realized because the young trees will not be able to mature. Another difficulty in disease control of Hevea is that the resistant genome seems to be a less dominant gene. SALB-resistant 3

plants were crossed with high-yield H. brasiliensis and their offspring are still susceptible to fungal attacks.14

Plant biochemists had been working on fungicides and improved application

techniques over the past few decades. Weekly application of fungicides were attempted

to control the spread of the fungi; however, young plants are difficult to spray without

considerable set-back in growth. Furthermore, Hevea’s wintering cycle involves

secondary leaf fall and refoliation after wintering. Matured Hevea plants are susceptible

to fungal attacks as well. At this time, there are no solution for this problem and led to

large losses of plantations; thus, inhibiting NR production on a commercial scale in

Central and South Americans.7 As Wade Davis commented in his book, "a single act of biological terrorism such as the introduction of fungal spores so small as to be readily concealed in a shoe could wipe out the plantations, shutting down production for at least a decade. It is difficult to think of any other raw material that is as vital and vulnerable."9

In 2005, approximately 21 million tons of rubber was produced and 42% of it (9

million tons) was NR.2, 15 In terms of quantity by type, NR is still the largest. (42% NR,

20% Styrene-butadiene rubber (SBR), 14% latex SBR, 12% polybutadiene, 5% ethylene propylene diene monomer rubber (EPDM), and 7% other synthetics.16) While the USA is

self-sufficient in synthetic rubber production, with substantial export activities, no Hevea

NR is produced in the USA. It is important to emphasize that no synthetic rubber matches

the performance of imported Hevea NR. The development of domestic supply of NR is

recognized in the Critical Agricultural Materials Act of 1984 (Laws 95-592 & 98-284).17,

18 The Act recognizes that NR latex is a commodity of vital importance to the economy

and the defense of the nation. Recently, due to the increased demand of NR from China, 4

Vietnam had doubled its production of NR from 155 thousand tonnes to 320 thousand tonnes in from 1995 to 2001.16 This caused a significant drop in NR market prices. In an attempt to control NR prices, the major NR producing countries (Thailand, Indonesia and

Malaysia) formed an International Tripartite Rubber Council (ITRC) to manage sales and stocks. Since the intervention of the ITRC, NR prices raised from ~20 cents US dollars

(USD) / kg to ~3 USD / kg of NR.

Scientists had been working on NR replacement since World War II (WWII). As a result, new polymers were synthesized and led to the development of SBR, polyvinyl chloride (PVC), chloroprene rubber (CR) and butyl rubber. The growth of synthetic rubbers (SR) was triggered mainly by geo-political events, when NR was in very short supply and high in demand. NR and SR share much of the processing technology; they are first prepared as un-vulcanized compounds and then vulcanized at higher temperature to give the final product.19 In short, the production cycle of both NR and SR is much

longer than thermoplastics due to vulcanization and mixing cycles. In addition, scraps

cannot be easily recycled.19

Despite SR share the same processing technology, their physical properties are not superior to NR particularly in green strength and low hysteresis. Green strength is the tensile strength and/or tensile modulus of an uncured rubber compound. This important processing property relates to the compound’s performance in extrusions, calendaring and tire building, particularly the 2nd stage building machine for radial tires. If tires are

constructed with rubber compounds with poor green strength, they may fail to hold air

during normal expansion in the 2nd stage of tire building prior to cure. While

5 polybutadiene is currently used to construct ~70% of the treads and sidewalls, the use of

NR in tires cannot be replaced or omitted.

In order to resolve these problems, thermoplastic elastomers (TPEs) were developed as a solution to reduce costs of production and processing of rubbers. EPDM blends with polypropylene (PP), thermoplastic polyurethanes, and polystyrene block copolymers blended with PP are a few examples of TPEs.20 Even though many new materials are introduced as NR replacement, the shift to thermoplastic technologies does not impact the NR industry to a great extent. Despite the cost efficiency, the thermoplastic replacement counterparts still do not always perform as well as NR in many ways.

This thesis will investigate NR biosynthesis to get a better understanding how the tree produces NR. Insight into the biosynthesis may lead to the development of a biomimetic system that would produce NR.

CHAPTER II

BACKGROUND

2.1. Chemical Structure of Natural Rubber

From the pyrolysis of NR, scientists realized that NR can be broken down to isoprene (IP). NR is stable below 200 oC and significant decomposition into smaller

fragments takes place beginning at 290-300 oC.21, 22 The main pyrolysis product of NR is

DL-limonene when pyrolysis is carried out below 450 oC (Figure 2.1.1a) and IP when further pyrolysis (retro Diels-Alder reaction) is performed (Figure 2.1.1b).21

b) a)

isoprene DL-limonene

Figure 2.1.1 - DL-limonene (a) and isoprene (b).

From this understanding, Bouchardat reported the first polyisoprene (PIP)

synthesis in 1879 using IP as the monomer and hydrochloric acid as the initiator.23-25 This

report quickly triggered research interests; however, the subsequent work was slow and not fully successful because high molecular weights were not reached and the mechanical

properties of the cured PIP were poor.24, 26 7

The how and why of rubber formation in Hevea trees are long-standing mysteries.

It was the analysis of Hevea rubber that led to the acknowledgement of the existence of macromolecules where the monomer units are connected by covalent bonds. Samuel

Shrowder Pickles of the Royal Institute of London was the first to propose a chain structure based on the building block27, 28 as shown in Figure 2.1.2a. Prior to 1910, NR

was believed to have a structure based on the self-assembly of dimers (Figure 2.1.2b)

held together by physical forces.28 Carl Harries based this proposal on the fact that he

found no end groups in natural rubber. Chemists at the time did not want to accept the existence of macromolecules.19, 29

Figure 2.1.2 - Suggested building blocks of NR before (b) and after (a) 1910.27

Infrared and nuclear magnetic resonance (NMR) spectroscopy, and X-ray studies

have shown that the major component of NR from Hevea brasiliensis is polyisoprene

(PIP) in cis-1,4 configuration, with about 6 wt% non-rubber components (mostly

proteins).30-33 The general agreement from the literature is that Hevea NR contains a

dimethyl-allyl head group followed by 2 units in trans-1,4 configuration from the

initiator, followed by >5000 cis-only units (Figure 2.1.3).33-35 Hevea NR has very broad

36, 37 molecular weight distribution (Mw/Mn ~ 2-15). Biochemical studies provide no information on the termination step of rubber formation; a variety of end groups

(hydroxyl, aldehyde, and amine, etc) have been found.23, 28, 29

Figure 2.1.3 - Chemical structure of NR.3 Head (α) and end group (ω) structure is not proven.

High molecular weight (MW) NR produced by other plant species such as

Parthenium argentatum (Guayule) and Ficus elastica have similar structures.34, 38, 39 13 C and 1H NMR spectroscopy are powerful tools for structural analysis of polymers; however, the exact structure of NR remains unproven due to its very high MW.33, 35

Figure 2.1.4 shows the 13C NMR spectra of PIPs isolated from Hevea and Guayule.33, 40

The signal at 32.3 ppm is assigned to carbon A (-CH2-C(CH3)=), and the signal at 26.4

ppm is assigned to carbon D (=CH-CH2-). The peak at 23.3 ppm is assigned to the carbon

of the methyl group (CH3-C(CH2)=) attached to carbon B. The signals at 135.1 and

125.1ppm are assigned to the olefin carbons B (=C(CH3)(CH2)-) and C (=CH-CH2-).

Figure 2.1.4 - 13C NMR spectra of NR from Hevea40 (a) and Guayule33 (b).

As mentioned before, the head and end group structures remain unproven. Tanaka analyzed naturally occurring lower MW (Mn ~ 20,000 g/mol) NR from a mushroom

(Lactarius volemus).33, 41 Figure 2.1.4 shows the 13C NMR spectrum with our assignments. The signal at 32 ppm is assigned to carbon A (-CH2-C(CH3)=), and the

signal at 27 ppm is assigned to carbon D (=CH-CH2-) in the main chain. The peak at 22

ppm is assigned to the methyl group carbon E (CH3-C(CH2)=) attached to carbon B. The

signals at 138 and 123 ppm are assigned to the olefin carbons B (=C(CH3)(CH2)-) and C

(=CH-CH2-). Tanaka assigned the signal at 18 ppm to the methyl carbons of the dimethyl

allyl head group (Fα), and the signals at 130 ppm to the olefin carbon Bα,tr1 of the trans

41 head group. The peak at 40 ppm was assigned to carbons Aα,tr1 and Aα,tr2 of the methylene groups of the two trans units attached to the dimethyl allyl head group (-CH2-

C(CH3)=). The methyl protons in the first two trans α-units (note that Tanaka labeled the head group as ω, so we changed the notation on the original NMR spectrum according to

polymer chemistry principles) were labeled as Eα,tr1 and Eα,tr2 and appear around 16 ppm.

As for the ω-end group, Tanaka assigned signals at 142 and 120 ppm to the olefin 10

’ carbons Bω (=C(CH3)(CH2)-) and Cω (=CH-CH2-) respectively. He also suggested the

41 peak at 60 ppm to be the methylene carbon next to an l ester end group (Gω). The carbonyl carbon (H) appears at 176 ppm, and signals I, J, K and L were assigned to the fatty acid residue of the ester end group. The suggested structure of Hevea NR is based on this work.

Figure 2.1.5 - 13C NMR spectrum of NR from L. volemus.33, 41 11

It is well-established that NR is composed of long-chain branched molecules.42-44

Fulton et al. showed the presence of long-chain branching and gel in Hevea NR by Field

Flow Fractionation (FFF): the log Rg – log MW conformational plot was shown to have a slope of 0.3 in the high MW range.45 Hevea NR contains 50–70% gel. Tanaka established that the gel has two different components that he termed “hard” and “soft” gel. The soft gel is produced by hydrogen bonding between phosphates and phospholipids at the ω-end group of Hevea NR. It has been shown that the addition of 1–2% ethanol into a Hevea

NR solution in a good solvent, such as toluene, dramatically decreases the gel content by breaking the hydrogen bonds that make up the “soft” gel, as shown in Figure 2.1.5.33, 37

Figure 2.1.6 - Deproteinization by treatment with 1~2% ethanol.37 (DMAPP denotes dimethylallyl pyrophosphate, FPP denotes Farnesyl pyrophosphate.)

The “hard gel” crosslinking points are believed to be formed by radical reaction between sulfur containing proteins and the dimethyl allyl double bond in the α-head groups of NR. These covalent bonds can be broken via transesterification.46

The exact structure of high MW NR remains to be proven by the next generation of scientists.

2.2.History of Natural Rubber

NR has a long and distinguished history. Ancient Mesoamerican peoples mixed the rubber latex harvested from the Castilla elastica shrub with the juice of Ipomoea alba

(a species of the morning glory vine) and produced solid rubber. Rubber balls made by the Olmec of Mesoamerica (often referred to as the “rubber people”) dated back to 1600-

1200 B.C.1 Mexican excavations found rubber balls dated 600 A.D. that were used in religious ceremonies and ball games. In 1403, Columbus reported that the natives of Haiti played games with balls prepared from the gum of a tree. In 1520, Fernando Cortes, a

Spanish “conquistador” observed rubber balls in games played at the Inca king

Montezuma’s palace.47 In 1735, Condamine, French explorer, brought a sample of rubber from Peru to France, made from the latex called “cahuchu” (milk of the tree) by natives.

He spelled the word as “caoutchouc” in French, which was later spelled as “kautschuk” in German.48 In 1752, Priestly coined the word “rubber”, referring to the ability of the material to “rub out pencil marks”.47 In 1820, McIntosh made a raincoat from cloth impregnated with NR. The coat was water repellent - in Great Britain the word

“McIntosh” is still synonymous with “raincoat” - but it was smelly and sticky.49 The

stickiness of rubber arises from free polymer chains moving independently. Hancock

perfected the process by “curing” or crosslinking the rubber using sulfur which made it

lose its stickiness because these free chains are now chemically bonded. As mentioned before and according to legend, Charles Goodyear accidentally dropped NR mixed with sulfur and white lead on a hot stovetop in 1839. This process cured NR fast enough to become the foundation of rubber technology.1 Akron, Ohio became the “rubber city”

when Seiberling and Miles built their first tire manufacturing plant and named it after

Goodyear. Buchtel College (today the University of Akron) offered the first course in

rubber chemistry in 1914.

World War II (WWII) threatened the access of the Allied Forces to the sources of

NR, by then mostly in Southeast Asia, because a pathogenic attack wiped out the

plantations in Brazil. Every tire, hose, seal, and valve, which were essential to war craft

required rubber. The “Synthetic Rubber Procurement Program” headquartered in Akron

was second in order of importance only to the “Manhattan Project” to make the atomic

bomb.50 The ultimate goal of the rubber program was to establish a domestic source of

rubber. This surge of research and commercialization effort led to the production of a

wide range of synthetic rubbers SR. In addition to the focus of trying to synthesize a NR

replacement, the program explored alternative plants to produce NR, including planting

Russian Dandelions in 41 states.51, 52 By 1964, SR made up 75% of the market53;

however, the ultimate goal of completely replacing NR with a synthetic alternative was

not accomplished.

The re-emergence of NR research was marked by the OPEC oil embargo of 1973,

which nearly doubled the price of SR. In addition, an unexpected threat was brought to

14 the synthetic market by the rapid adoption of the radial tire. The construction of radial tires necessitates the use of NR that possesses the required physical properties. By 1993, natural rubber had recaptured 39% of the domestic market.53 Today we still do not have a synthetic replacement.

In the next Chapter we will briefly review the history of efforts to produce an NR equivalent synthetically.

2.2.1. Synthetic Polyisoprenes

It has long been a fascination of chemists to create a chemical and physical equivalent of NR. Various catalytic systems have been investigated.54-57 Table 2.2.1 summarizes the microstructures of synthetic polyisoprenes (PIPs) obtained by various polymerization techniques. The possible microstructures of PIP are shown in Figure

2.2.1. It can be seen that no methods produced fully cis-1,4 PIP enchainment.

Table 2.2.1 - Microstructures in synthetic PIPs

Figure 2.2.1 – PIP microstructures. *an example of C15 is shown, other cyclics may occur.

The cationic polymerization of IP was studied first by Gaylord et al. in 1966.26 It was found that AlEtCl2 or SnCl4 in n-heptane yielded low conversion (~20%) and the resulting PIP mostly consisted of trans-1,4 chain enchainment with a few repeat units

26 with cis-configuration. TiCl4 in n-heptane was found to be inactive unless hydrous n- heptane was added. The rates of polymerization in aromatic solvents were generally much higher due to extensive chain transfer to solvent. The stereo-regularity of the resulting polymers was found to be mostly trans-1,4 with Mns around ~50,000-100,000

58 g/mol and Mw/Mn ~8 due to cyclic side products. In halogenated solvents, the rate of polymerization was reported to be the fastest among the three types of solvents discussed.

In solvents of higher polarity, such as o-dichlorobenzene, more linear structures were obtained. The resulting polymers contained higher cis-content (~30%).58 In a more recent study in 2008, Khachaturov et al. initiated cationic polymerization of IP by a TiCl4- trichloroacetic acid system in dichloromethane.59 They reported ~50% cyclized product and ~50% loss of unsaturation due to cyclization. Of the remaining unsaturation ~47% was found to be trans-1,4, with ~1.5% cis-1,4 and ~1.5% 3,4.60

The radical polymerization of IP was also investigated in 1960s. Gobran et al.

studied the bulk polymerization of IP using 2-azo-bisisobutyronitrile (ABIN) and benzoyl

o 61 peroxide as initiators at 60~90 C. The authors obtained low MW PIPs (Mn =

1,000~7,000 g/mol) and determined that the radical polymerization of IP was a dead-end polymerization. Dead-end polymerization is a term coined in the 1950s by Tobolsky

describing a polymerization that can only reach a limiting conversion due to initiator

depletion.62 In the 1990s, Solomon, Moad and Rizzardo showed the first examples of nitroxide-mediated free radical polymerization (NMP) of vinyl monomers.63 Hawker et al. attempted the NMP (Nitroxide Mediated Radical Polymerization) of IP using 2,2,5- trimethyl-4-phenyl-3-azahexane-3oxy-nitroxide.64 (Figure 2.2.2) In 36 hours, they

64 achieved 75% conversion with Mn = 19,800 g/mol and Mw/Mn = 1.07.

Figure 2.2.2 – NMP of IP using 2,2,5-trimethyl-4-phenyl-3-azahexane-3oxy-nitroxide as initiator.64

The authors did not explore the microstructure of the resulting polymer. In 2006,

Perrier et al. attempted controlled radical polymerization of IP via reversible addition- fragmentation chain transfer (RAFT).65 The authors used two different RAFT agents: 1)

2-(2-cyano-propyl)dithiobenzoate and 2) 2-ethylsulfanylthiocarbonyl-sulfanylpropionic

65 acid ethyl ester (ETSPE). They were able to achieve 97.2% conversion, Mn = 27,400 g/mol, and Mw/Mn = 1.47 in 72 hours using the ETSPE RAFT agent. The resulting 17

polymers had 75% 1,4-, 20% 3,4-and 5% 1,2-addition.65 Atom transfer radical

polymerization (ATRP) of IP have not been realized because dienes chelate to the copper

catalyst.65

The anionic homopolymerization of IP was researched extensively in the 1950s

and 1960s. Tobolsky et al. did a systematic study of IP polymerization with various

solvent systems.66, 67 The authors were able to obtain 94% cis-1,4 polyisoprene initiated

66 with n-butyllithium in n-heptane with Mυ = 2,000~150,000 g/mol. They concluded that

high cis-content can be obtained by using hydrocarbons as the solvent.67 Morton et al.

studied the effect of addition of tetrahydrofuran (THF) into the anionic polymerization of

IP in 1963.68 The authors found that 3,4-enchainment increased dramatically.68, 69

Bywater et al. continued this study using cyclohexane/THF solvent systems and found that the polarity of the solvent affected the effective counter cation size.70 The recent

literature on anionic polymerization of IP has focused on block copolymer synthesis.71-73

Ziegler-Natta and metallocene catalysts yielded the highest cis content in IP polymerizations. More recently research focused on Nd-based catalysts activated by R3Al that display high cis-1,4 stereo-specificity in both the homo- and copolymerization of

IP.74 However, early versions of these catalysts were limited by the lack of control over

56 MWs and Mw/Mns. It was hypothesized that this problem was attributable to the

heterogeneity of the catalyst systems and the multiple active species.56 In 2003, Dong et

al. circumvented these problems by introducing a [Nd(O-iPr)3]/modified

methylaluminoxane (MAO) catalyst system.75 The improved solubility of MAO in

heptane allowed for a homogeneous single active site system. The authors have tried

various reaction conditions for this system. In heptanes an [Al]:[Nd] ratio of 100:1 at 30

4 °C gave 98.9% conversion with Mn ~5x10 g/mol, Mw/Mn = 1.14 and 91.4% cis-1,4

structure.75 In toluene, an [Al]:[Nd] ratio of 300:1 at 30 °C gave 100% conversion with

4 75 91.7% cis-1,4 structure with Mn ~8x10 g/mol and Mw/Mn = 2.21. In dichloromethane,

the authors obtained mainly cyclized trans-1,4 structures. In their 2005 follow-up study

of Nd-based metallocene, Taniguchi and Dong et al. reported a ternary catalyst system

composed of Nd(III) isopropoxide, dimethylphenylammonium

tetrakis(pentafluorophenyl)borate, and triisobutylaluminum.56 They found the optimal

catalyst composition to be a [Nd]:[borate]:[Al] = 1:1:30, which gave greater than 97%

conversion, Mn = 200,000 g/mol, Mw/Mn of 2.0, and approximately ~90% cis-1,4

structure.56 Unfortunately these systems yielded much lower MWs than that found in

Hevea NR. The closest commercial attempt to mimic NR was accomplished by the

Goodyear Tire&Rubber Company with their product called Natsyn® produced with a

5 titanium-aluminum (Ziegler-Natta type) catalyst with Mn = ~2x10 g/mol, Mw/Mn of ~3 and ~98.5% cis content. 55

It is our belief that in order to be able to create SR that mimics the structure and

properties of NR, we need to understand NR biosynthesis. The following sections

summarize our understanding of in vivo and in vitro NR biosynthesis from the viewpoint

of polymer chemistry.

2.3. Natural Rubber Biosynthesis

2.3.1. Rubber Producing Plants

NR is obtained from latex, an aqueous emulsion present in the laticiferous vessels

(ducts) or parenchymal (single) cells of rubber-producing plants (Figure 2.3.1). Although

more than 2,500 plant species are known to produce NR, there is only one important commercial source, Hevea brasiliensis (the Brazilian rubber tree). The rubber

from Guayule, Parthenium argentatum, is being developed as a non-allergenic NR

mainly for rubber gloves.76, 77

The rubber latex from H. brasiliensis is harvested by “tapping” the rubber tree.

An incision is made on the trunk and latex will ooze out of the incision. This white liquid

is collected and then coagulated to yield high molecular weight (Mn >1 million g/mol) polymer. Guayule, Parthenium argentatum, is grown in the Southwestern USA. Guayule latex is mainly used for advanced medical and consumer products. The guayule rubber is produced by a green aqueous-based extraction process on a commercial scale.78 Russian dandelion, Taraxacum kok-saghyz, is being developed as a rubber-producing crop in the

Northern USA.38, 79 Figure 2.3.1 shows an array or rubber producing plants that have

been of research and commercial interest. Hevea is the most studied NR, so Hevea will

be discussed in more detail.

Figure 2.3.1 - Examples of rubber producing plants.

2.3.2 Anatomy of the Hevea NR Latex

The productivity of Hevea trees is usually as high as 50–100 g latex per day in a mature tree.80 Depending on the seasonal effects and the state of the soil, the average

composition of Hevea latex is 25~35 wt% cis-1,4-polyisoprene (PIP), 1~1.8 wt% protein;

1~2 wt% carbohydrates; 0.4~1.1 wt% lipids, 0.5~0.8 wt% amino acids, and 50~70 wt%

water.81 The rubber particles have a distribution of diameters ranging from 0.1 to 10 μm33

(Figure 2.3.2a) and are stabilized in the cytosol by a membrane of phospholipid monolayer (Figure 2.3.2b).4 The cytosol contains mainly water, salts, organic molecules and ribosomes.82 The cis-prenyltransferase enzyme (Figure 2.3.2b) is a membrane-bound amphiphilic enzyme found in NR producing plants, yet to be isolated and fully characterized.33, 83

Figure 2.3.2 - Visualization of NR particles and its structure.18

It has been known since the 1950s that the “chain elongation” of rubber molecules proceeds by the addition of the isopentenyl pyrophosphate (IPP) monomer.84, 85 The initiator is believed to be farnesyl pyrophosphate (FPP).86 These molecules are water-

21 soluble and are present within the cytosol along with the divalent metal “cofactors”

(Figure 2.3.2b), needed to be present in the biosynthesis process that will be discussed in

the next section. The biosynthesis of NR is catalyzed by the rubber transferase enzyme, cis-prenyl transferase.4, 84, 87, 88 Enzymes are proteins that act as highly selective catalysts

in biochemical reactions. The rubber transferase is integrated into the phospholipid

monolayer that surrounds the latex particles. The phospholipid monolayer stabilizes the

particles to prevent aggregation in the aqueous medium. The hydrophobic polymer chains

reside within the latex particles, and polymerization proceeds at the active sites of the

enzyme.34 The rubber transferase enzyme, which has not been isolated yet, is proposed

to be amphiphilic, with hydrophilic regions facing the cytosol to allow the access of

hydrophilic building blocks and the hydrophobic regions accommodating the growth of

NR.

2.3.3 Biochemical Pathway of Natural Rubber Biosynthesis In Vivo

As mentioned before, it is accepted in the literature that the monomer to make NR

is IPP. Figure 2.3.3 shows the structure of IPP, which can be considered to be an adduct

of pyrophosphoric acid (H4P2O7) and isoprene (IP).

Figure 2.3.3 - Structure of isopentenyl pyrophosphate IPP at pH = 7.4.89

At the physiological pH of 7.4, IPP is a stable di-anion with two potassium

counter cations. The cytosol has a high concentration of potassium ions (139 mM) and low concentration of sodium ions (12 mM) for osmoregulation purposes.82 In contrast, in

the human blood, the potassium concentration is only 4 mM and the sodium

concentration is145 mM.90

IPP is produced from carbohydrates in plants, bacteria, algae, and mammals,

including humans. The synthesis of IPP can proceed by two different pathways: the

mevalonate (MVA) pathway and non-mevalonate (non-MVA) or deoxy-xylulose

pathway. These two distinct pathways have evolved in different organisms. In eukaryotes

(cells that contain nuclei that carry genetic information), IPP is derived from acetyl-CoA

(coenzyme A), while in prokaryotes (cells that lack nuclei) and plant chloroplasts, IPP is

derived from 1-deoxy-D-xylulose-5-phosphate.91 In higher plants, the mevalonate pathway operates mainly in the cytoplasm and mitochondria, whereas the non-MVA

pathway operates in the plastids.92 Plastids are sub-units in a plant cell that are

responsible for photosynthesis; they also serve as storage units for fatty acids and terpenes found in plants. It is suspected that the IPP for NR biosynthesis is derived from the MVA pathway located in the cytosol.93 However, IPP produced by the 1-deoxy-D-

xylulose-5-phosphate pathway may also diffuse from the plastids to the cytosol.94

The biosynthesis of polyisoprenoids or terpenoids is believed to be regulated by a series of chemical equilibria shown in Figure 2.3.4. IPP is first isomerized into dimethyl allyl pyrophosphate (DMAPP) by the isomerase enzyme. DMAPP may serve as the initiator for subsequent terpenoid synthesis (chain growth or polymerization) catalyzed by appropriate enzymes. Longer chain initiators (Figure 2.3.5) are synthesized by trans-

23 prenyltransferases. When excess IPP is present, the equilibrium shifts toward DMAPP which in turn can be converted into isoprene (IP) by the isoprene synthase enzyme.Because of its low boiling point (34oC) IP easily evaporates, so the corresponding equilibrium is shifted towards IP to remove excess IPP. The estimated rate of IP production by the human body is 0.15 µmol/kg/h, which corresponds to about 17 mg each day for a 70 kg person. Vegetation emits 600 megatons of IP per year into the atmosphere – major producers are oak trees, tropical broad leaf trees and shrubs. Figure

2.3.4 presents our rendition of the IP biocycle.

Figure 2.3.4 - Terpenoids biosynthesis cycle.

The C15 farnesyl pyrophosphate (FPP) shown in Figure 2.3.5 is believed to be the initiator in the in vivo biosynthesis of Hevea rubber based on numerous reports of α-trans

head groups found in 13C NMR spectrum of low MW rubbers, but it is not proven for

Hevea.33

For better chemical understanding of the compounds, we will use –OPP to denote pyrophosphate groups. In the biological literature, the pyrophosphate group is typically written as –PPi. In the presence of the metal cofactors (Mg2+ or Mn2+ in vivo and Mg2+ in vitro) and the rubber transferase enzyme, an IPP unit adds to extend the FPP by one more unit but in cis configuration. Each step is accompanied by the liberation of pyrophosphoric acid. This step is repeated and the process generates NR with more than

5000 units. (Figure 2.3.5).

+ OPP OPP

Farnesyl pyrophosphate Isopentenyl pyrophosphate

cis-prenyltransferase metal2+ cofactor

OPP HOPP 2

cis-prenyltransferase metal2+ cofactor

OPP 2 >5000

HOPP

Figure 2.3.5 - Synthesis of NR in H. brasiliensis. OPP stands for the pyrophosphate end- group and HOPP represents pyrophosphoric acid.

The FPP initiator has successfully been used to make Hevea rubber in a culture tube (in vitro biosynthesis, to be discussed later). Other oligomeric allylic pyrophosphates

(geranyl-, farnesyl- and geranyl-geranyl-pyrophosphate) can also serve as initiators in vitro. The chemical structures are shown in Figure 2.3.6.

Figure 2.3.6 - Structure of the allylic oligoisoprene pyrophosphates. DMAPP: dimethyl allyl pyrophosphate, GPP: geranyl pyrophosphate, FPP: farnesyl pyrophosphate, GGPP: geranylgeranyl pyrophosphate.

Our rendering of the initiation and propagation steps are shown in Figure 2.3.7 using symbolism from biochemistry. When compared with the polymer chemistry symbolism shown in Figure 2.3.5, both show the stepwise addition of the building blocks, with the elimination of a pyrophosphoric acid in each step. The monomer (IPP) is termed

“substrate” while the initiators are termed “cosubstrate” in the biochemical literature.

Figure 2.3.7 - Biochemical representation of rubber biosynthesis.34

As mentioned before, enzymatic activity requires the presence of divalent cations such as Mg2+ or Mn2+ in vivo , called “activity cofactors”.4 The exact role of the cofactors is still unclear. Termination is believed to occur when the chain end detaches from the enzyme. During this step the –PP terminus may hydrolyze, yielding an -OH end group as shown in Figure 2.1.3. This end group may react with fatty acids, yielding ester end groups shown in Figure 2.1.4.

2.3.4 Prenyltransferases

Prenyltransferases are enzymes that catalyze the synthesis of various isoprenoid compounds: sterols, terpenes, and natural rubber. Based on the configuration of the repeat

units in the products, prenyltansferases are classified into two classes: trans- and cis- prenyltransferases. In both prokaryotes and eukaryotes, trans-prenyltransferases catalyze the formation of geranyl diphosphate (GPP:C10), farnesyl diphosphate (FPP:C15), and geranylgeranyl diphosphate (GGPP:C20) (Figure 2.3.6) These compounds then serve as initiating species to produce many other longer chain isoprenoids necessary for cellular growth and survival. Figure 2.3.8 shows the structure of an avian trans-prenyltransferase that catalyses FPP synthesis from DMAPP and IPP. The binding sites for the DMAPP and IPP were located within the hydrophilic regions of the enzyme (red arrows), whereas chain growth was proposed to take place within a hydrophobic pocket positioned toward the bottom end of the conical enzyme (Figure 2.3.8).

Figure 2.3.8 - A scheme of the active sites in an avian trans-prenyltransferase.95 The oval represents a large motive proposed to stop chain growth.

Mutational analyses and x-ray crystallographic investigations revealed crucial amino acid residues in the conserved domains of various trans-prenyltransferases to stop

the chain growth, accounting for the mechanism of chain length determination.95-97 The

structural genes for FPP synthase98-100 and GGPP synthase101-103 have been cloned and

characterized from various organisms. However, only three cis-prenyltransferase genes

have recently been cloned104, 105 from E. coli, M. luteus and yeast, which share a low level of sequence homology (~30%). The genetic sequences of the rubber transferase remain unidentified because it is a membrane-bound enzyme in low abundance.4 The fact that the enzyme activity is rapidly lost upon disruption of the structural integrity of the membrane poses one of the great challenges in sequencing the enzyme.87, 106 Generally, rubber

transferase is accepted as an amphiphilic enzyme residing at the interface of the latex

(Figure 2.3.8). Propagation occurs when the initiator or the NR molecule is bound at the specific catalytic site within the funnel-like crevice of the enzyme and is activated by co-

factors, and IPP enters from the aqueous phase.

2.3.5 Mechanism of Prenylation in Short Chain Isoprenoids

The mechanism of short-chain terpenoid biosynthesis proposed and proven by

Poulter86 is illustrated in Figure 2.3.9. According to the authors, resonance stabilized allylic carbocations are generated by the dissociation of the –OPP end group. Then the allylic carbocation reacts with the double bond of the IPP, and a pyrophosphoric acid,

HOPP, is released. The dissociative mechanism (SN1) was proven by constructing the

Hammet plot for prenyl transfers with fluorine substituted allylic pyrophosphates.

OPP prenyltransferase OPP

OPP OPP

HOPP H OPP

Figure 2.3.9 - The mechanism of terpenoid biosynthesis proven by Poulter et al.86

The polymer chemistry community has been reluctant to accept this mechanism,

although ionic polymerizations in aqueous media have been demonstrated.107

2.3.6 Proposed Mechanism of Natural Rubber Biosynthesis: Natural Living

Carbocationic Polymerization (NLCP)

Based on Poulter’s work, Puskas et al. proposed that NR biosynthesis most likely

also proceeds by a carbocationic mechanism (Figure 2.3.10).35 According to this

proposal, the initiation starts with an enzyme (and divalent cofactor) – assisted ionization

of the carbon-oxygen bond of the initiator and yields an allylic carbocation plus a

pyrophosphate counter-anion; the enzyme plus cofactor(s) coordinate with the pyrophosphate “protecting” group and mediate the formation of the initiating

carbocation. According to polymer chemical convention, the enzyme plus cofactors

constitute the co-initiating system. Ionization at the chain end is favored by resonance

stabilization of the allylic carbocation and increasing entropy of the system.

Subsequently, IPP adds to the allylic carbocation, yielding a tertiary carbocation which,

via proton elimination, regenerates the trisubstituted allylic pyrophosphate.35 This

mechanism applies to the formation of trans-1,1-dimethylallylic initiators (natural initiators invariably are trans), catalyzed by trans-prenyl transferase, as well as to the incorporation of the subsequent cis-units catalyzed by cis-prenyl transferase. In regards to trans or cis-stereoregulation, Puskas proposed that the specific enzyme functions as the template. The template theory was originally proposed by McMullen108. He suggested

that polynucleotides complex with the rubber and act as template. However, it has been

demonstrated that polynucleotides are not involved in NR biosysthesis. The incorporation

of each IPP unit is always accompanied by the loss of pyrophosphoric acid HOPP (or its

salts).

Figure 2.3.10 - Proposed NLCP mechanism of NR biosynthesis.35 31

This mechanism is very similar to Yokozawa et al.’s109-111 general mechanism for the biosynthesis of many natural biopolymers. These authors developed the concept of

“chain-growth polycondensation”, according to which an enzyme activates the initiating entity and/or the dormant polymer chain end, which proceeds to add monomer and the enzyme complex relinquishes the protective end group to regenerate the chain end.

Figure 2.3.11 helps to visualize Yokozawa’s concept. As opposed to synthetic polycondensation, the monomer is inactive and can only react with the activated initiator or polymer chain end.

Figure 2.3.11 - Yokozawa’s concept of chain-growth polycondensation.109, 110

Examples cited by these authors include peptide extension (termed “elongation”

in biochemistry), DNA and RNA syntheses and natural rubber biosynthesis.109-111

Yokozawa’s group developed two strategies for chain-growth polycondensation. The first

32 strategy involved the activation of polymer end groups, and led to aromatic polyamides, polyesters, polyethers, poly(ether sulfone) and polythiophene.110 By the second strategy, the monomer was separated from the polymerization phase to prevent monomer- monomer and polymer-polymer condensations. Yokozawa’s work was a breakthrough in biomimetic polymer synthesis and produced living polycondensation reactions displaying narrow MWDs. The synthesis of cis-PIP was attempted as shown in Figure 2.3.12. The electrophilic initiator (triphenylmethyl (trityl) perchlorate) was postulated to create an allylic end group structure. This structure was expected to be reactive with the unreactive monomer structure, accompanied by the elimination of the protecting allyl trimethyl silane group. However, only elimination happened and a diene was obtained.110

Figure 2.3.12 - Attempted “bio-inspired” synthesis of cis*-1,4-polyisoprene.110 *(The resulting polymer was in the trans conformation)

The proposed chain-growth polycondensation mechanism should lead to monodisperse PIP; however, all natural rubbers exhibit multimodal/broad molecular weight distributions. This most likely is due to continuous initiation with simultaneous

chain growth. It should be noted that all industrial rubbers exhibit broad MWD which is

required for better processing and the required balance of properties.

2.4 In Vitro Natural Rubber Biosynthesis

The first in vitro polymerization system was described by Archer et al. in the

1980s. They incubated 14C-IPP in the presence of unlabeled neryl pyrophosphate (NPP)

or geranyl pyrophosphate (GPP) initiators in a suspension of Washed Rubber Particles

(WRP) isolated from living Hevea latex, and showed the incorporation of the

radiolabeled IPP.84 The presently accepted method to determine the activity of rubber

transferases is based on this work: the activity is calculated based on the incorporation

rate of 14C IPP monomer into existing chains or forming new chains from the initiator in

vitro.87, 112, 113 This is accomplished by establishing a double reciprocal plot of kinetic

data (a.k.a. Lineweaver-Burk plot) with 14C-IPP. The isolated WRPs are incubated with a

predetermined amount of APP (Allylic Pyrophosphate) initiator and 14C-IPP (substrate)

for 60 minutes. The unreacted 14C-IPP monomers were extracted from the rubber and the

radioactivity was measured by scintillation spectrometry (SS). In enzyme kinetics,

biochemists report the enzymatic activity by reporting the Michaelis-Menten (binding)

114, 115 constant (KM) of an enzyme. To determine this value, biochemists flood the enzyme

with substrates to a limiting condition where all the active sites are bound with substrates

and reach a theoretical maximum reaction velocity (VMax). At this point, the reaction rate

of the enzyme is limited to the intrinsic turnover rate of the active site. The inverse of the

reaction velocity data is plotted against substrate concentration to determine the

Michaelis-Menten constant (KM), which can be found from the x-axis intercept of the

114, 115 Lineweaver-Burk plot. (Figure 2.4.1). The slope of the plot is KM/Vmax.

Figure 2.4.1 – Lineweaver-Burk Plot of inverse reaction velocity (V) vs IPP concentration in the determination of the enzymatic activity of Guayule WRP.116

By comparing the KM between WRPs from various plant species or even within batches of WRPs from the same plant, the enzyme activity of WRPs can be established. It was found that the rate of rubber biosynthesis in vitro increases with the size of allylic diphosphate initiator, and that initiation regulates the overall rate of rubber biosynthesis.

Therefore, FPP and longer isoprenoids are utilized as synthetic initiators for in vitro NR

biosynthesis systems.

Tanaka’s group established a new method for in vitro rubber biosynthesis using the fresh bottom fraction (BF) of NR latex.117, 118 After filtering out particles and coagulated rubber using a muslin cloth, the liquid latex was centrifuged. After centrifuging three phases were observed: an upper phase, a middle clear phase called C- serum (CS) and a bottom phase (BF).117 Instead of using the WRPs (top layer), the

35 authors used BF for in vitro NR biosynthesis as they believed that the BF contained all the necessary components for NR biosynthesis in comparison to the WRP. Using gravimetric analysis they observed that more than ~10 wt% new rubber was formed with the addition of 14C labeled IPP or FPP to fresh BF.118 The formation of new rubber was confirmed by the incorporation of 14C radioactive IPP into the resulting rubber.118 Figure

2.4.2 compares the UV traces of the endogenous NR from BF and the in vitro NR rubber.

The BF has a high molecular weight fraction around ~106 g/mol with a lower MW tail.

Newly-formed rubber produced a peak at about ~105 g/mol. The radioactive traces also revealed that while the 14C-IPP incorporated into new chains, it also added to pre- existing chains in the lower MW tail fraction of the BF.118

Figure 2.4.2 - SEC trace of Hevea BF (dashed line) and in vitro NR (solid line) .118

It was also found that a small amount of new rubber formed in active BF without the addition of IPP and FPP, concluding that BF contains all of the enzymes and precursors necessary to produce rubber.119 Wititsuwannakul’s group in Thailand also 36

developed their unique Washed Bottom-fraction Particles (WBP)s for in vitro NR

biosynthesis.83 WBPs contains BF and non-rubber particulates such as lutoids

(membrane-bound bodies that cause flocculation of rubber particles) and Frey-Wyssling complexes (globules associated with plant metabolism).83, 120 It was observed that WBPs

are able to incorporate radioactive IPP into NR and the activity is enhanced in the

presence of sodium dodecyl sulfate detergent.120 This is most likely due to the fact that

detergents aid latex stabilization. More recently, Wititsuwannakul’s group investigated

the addition of two cloned suspected gene sequences of Hevea rubber transferase

expressed in E. Coli.83 It was found that a combination of one of the cloned Hevea rubber

transferase sequences with WBP promoted NR growth. In addition, the cloned Hevea

rubber transferase sequence by itself was able to synthesize polyisoprenoids chains in the

104 g/mol range.83

Benedict et al. investigated the in vitro synthesis of guayule rubber using WRP,

synthetic IPP monomer and DMAPP initiator.113 Their WRP was prepared from stems of

P. argentatum, after removal of the bark and impurities. The rubber latex was centrifuged and the top rubber particulate layer was collected and purified as the WRP. The authors used a SEC coupled with a scintillation spectrometer to demonstrate that radioactive IPP incorporated into NR.113 They observed that rubber was formed with a peak at ~105 g/mol within 15 minutes, and that the rubber was able to grow to ~106 g/mol in 180 minutes

(Figure 2.4.3).113

Figure 2.4.3 - in vitro guayule NR biosynthesis.113

Cornish et al. developed a method to produce WRPs that will be utilized in our studies. This method is an improvement of Benedict’s group’s WRPs preparation method, where the top rubber particles are collected and purified by repeated buffer washes and centrifugation.87 Her group determined the MW of in vitro NR by means of dual-labeled liquid SS.121 By introducing both radioactive IPP monomer and FPP initiator

into in vitro NR biosynthesis, an average MW of newly formed rubber could be calculated from a ratio between the 14C-labeled monomer and the 3H-labeled initiator.121,

122 The authors assumed that one initiator molecule is used to initiate one rubber chain,

and the IPP monomer is not consumed by the extension of pre-existing chains. Then, the

MW can be calculated as: [((14C-IPP incorporation rate) + 3(3H-FPP incorporation rate)) x MW of IP / [3H-FPP incorporation rate + diphosphate MW], which represents a

monomer / initiator ratio to obtain MW. The three in the formula represents the 3

isoprene units of FPP.121 This method works well in the 104-105 Da range. Outside of this range, the radioactivity of the initiator becomes very low and leads to experimental error. 38

Cornish et al. demonstrated that increasing the amount of IPP while keeping the FPP

concentration constant resulted in increased MW in three different types of WRPs

(Figure 2.4.4).121, 122 Cornish et al. published MW values which were lower than those found by Tanaka’s and Wititsuwannakul’s group where MWs were determined by SEC and SS signal from 14C-IPP. However, her findings support the notion of a living-like

polymerization.

Figure 2.4.4 - Experimental MW results from Cornish et al.’s in vitro NR system for three types of WRPs (Fig tree, Guayule, and Hevea) a) 0.25 µM FPP, b) 2.5 µM FPP.122

Her group studied molecular weight regulation using 14C-labeled monomer and

3H-labeled initiator in vitro with WRPs from Hevea, guayule and Ficus elastica.122 Using dual-labeled liquid scintillation spectrometry they showed that the latter produced twice as high MW NR in vitro than in vivo. Hevea and guayule WRPs, on the other hand,

produced much lower MW NR in vitro than in vivo. Thus MW regulation in NR biosynthesis remains unknown.

In summary, NR can be produced in vitro. However, the very complicated

process to isolate the active rubber transferase and synthesize the monomer renders this

process commercially not viable. Therefore it would be highly desirable to devise a

biomimetic process to produce an NR equivalent. Today, NR remains irreplaceable in

many important applications.

2.5 In Vitro Biosynthesis Using Modified Synthetic Initiators

Early studies of prenyltransferases often involved 32P-labeled isoprenoid initiators, given the ease of synthesis of these compounds.123, 124 However, since 32P was readily lost due to the prenylation mechanism and labile nature of the pyrophosphate group, the use of such compounds provided mixed results. Subsequently researchers tried to use traceable modified synthetic initiators. Indeed, prenyltransferases were found to be able to recognize some modified synthetic initiators. In 1985 Baba et al. utilized (E,E)-

2diazo-3-trifluoropropionyloxy geranyl pyrophosphate (DATFP-GPP, Figure 2.5.1) to isolate and identify a 30,000 Da protein subunit to be the binding site for the isoprenoid initiators in undecaprenyl pyrophosphate synthase.125 The authors determined the binding

efficiency by adding synthetic GPP and 14C-labeled IPP to in vitro NR biosynthesis.125

They have found that the binding efficiency for DATFP-GPP + 14C-labeled IPP was

~54% of that of the GPP + 14C IPP control experiment.125 The synthetic initiator with the

shorter C5 DMAPP allylic pyrophosphate (DATFP-DMAPP) was not able to bind to the

active site as no 14C IPP was found to be incorporated.125

O 3H

F3C OPP O

Figure 2.5.1 – Chemical structure of DATFP-GPP

While the DATFP-analogues accurately mimicked FPP, they suffered from low affinity and required prolonged short wavelength UV irradiation for photoactivation.

Distefano and Cornish et al. later developed a number of analogues of farnesyl and geranylgeranyl diphosphates containing a benzophenone chromophore for photoaffinity purposes.106 McMahan et al. then studied the benzophenone-modified diphosphate

analogues in three rubber-producing WRPs (Guayule, Hevea, and Fig tree).126 The ketone

group in benzophenone undergoes C-H bond insertion reaction upon excitation with 350

nm wavelength light. This covalently attaches the tracer to a variety of functional groups

present in the enzyme. The benzophenone-modified-FPP was observed to bind to the

active site of prenyltransferases. DMAPP analogues were less reactive, and GPP-

analogues were the least reactive.106

In this thesis we will present the synthesis of a modified initiator di-isobutylene-

neryl pyrophosphate (Figure 2.5.2). This is the model of a polyisobutylene-farnesyl

pyrophosphate macroinitiator that the Puskas group plans to investigate in NR

biosynthesis.

O O O O OPP Nerol-PP-DVA-TMP

Figure 2.5.2 – Chemical structure of di-isobutylene-neryl pyrophosphate.

2.6. Biomimetic Polymerization of IP

Biomimicry (from bios, meaning life, and mimesis, meaning to imitate) is a new

discipline that studies nature's best ideas and then imitates these designs and processes to

solve human problems.127 The core idea is that nature, imaginative by necessity, has

already solved many of the problems we are grappling with. Animals, plants, and

microbes are the consummate engineers. They have found what works, what is

appropriate, and most important, what lasts here on Earth. After 3.8 billion years of

research and development, failures are fossils, and what surrounds us is the secret to survival.

Based on the new understanding of NR biosynthesis Puskas et al. investigated the

cationic polymerization of isoprene (IP) initiated by allylic cations generated through

128 ionization of dimethylallyl bromide (DMABr) by TiCl4. (Figure 2.6.1) Using this

strategy, 1,4-oligoisoprene carrying a dimethyl allyl head group was produced in both cis- and trans- configurations, together with cyclized sequences.

Figure 2.6.1 – Biomimetic initiation of IP polymerization from Puskas et al.128

In a 2009 follow up study by Puskas and Deffieux et al., DMABr was replaced with dimethyl allyl alcohol (DMAOH) as the initiator.129 The authors attempted the polymerization in bulk, methylcyclohexane, hexanes, and dichloromethane at -40 to 20

oC. The resulting polymers were in oligomers with MWs of 500~1300 g/mol.129 The unsaturation retained in the oligomers was approximately 55%.129 The authors found that high IP conversions were found in less polar media and chain transfers occurred as Mns decreased without the presence of DtBP (2,6-di-tert-butyl pyridine) proton trap. NMR spectra showed that the unsaturation were mostly in trans-1,4 configuration.129

The polymerization of DMAOH was later studied with B(C6F5)3 as the co-

129 initiator in 2011. B(C6F5)3 used in this study is a Lewis acid (LA) with relatively inert

B-C bonds. Many boric LA decompose with the formation of B-F bonds. The authors initiated DMAOH with B(C6F5)3 and studied the resulting compounds. The authors found that the presence of DtBP affected the polymerization. In one case, the LA successfully cationized DMAOH for polymerization; however, it is also possible that DtBP led to elimination to generate IP monomer in situ to result in oligomerization.129

Puskas and Deffieux et al. then investigated the carbocationic polymerization of

IPOH (3-methyl-3-buten-1-ol) initiated by dimethyl allyl alcohol with BF3·2H2O as the co-initiator.130 (Figure 2.6.2). This chemical scheme was a synthetic analogue of NLCP described in Chapter 2.3.6. The pyrophosphate end group of NLCP was replaced by an 43 alcohol group. The polymerization proceeded to polymerize IPOH to yield oligomers

~1000 g/mol with Mw/Mn ~1.3. The structure of the product was quite complex and

MALDI-ToF mass spectrometry (MS) revealed that the reaction initially proceeded by

1,2-addition of monomer followed by chain transfer .130

Figure 2.6.2 – Possible carbocationic polymerization pathways for IPOH.129

In 2011, the cationic polymerization of IP using the 1-(4-methoxyphenyl)ethanol

/B(C6F5)3 initiating system in solution (dichloromethane or α,α,α-trifluorotoluene) and in

aqueous media (suspension, dispersion, or emulsion) was investigated by Puskas and

Deffieux et al.131 The authors observed that the reaction proceeded by controlled initiation, followed by irreversible termination in organic solvents.131 This resulted in polymers with Mn ~ 5000 g/mol, Mw/Mn ~2.5, and high content of intact double bonds

(~70%) in the polymer backbone. When using α,α,α-trifluorotoluene as the solvent, the

synthesis of polyisoprene resulted in chains with Mw/Mn ~1.4 and higher content of intact

double bonds (72~88%).131 When the polymerization was initiated with a trace amount of

water, polyisoprenes with fairly high molar mass (Mn ~18,000 g/mol) and Mw/Mn <2.4

were obtained. In aqueous media, the cationic polymerization of isoprene with 1-(4- methoxyphenyl)ethanol/B(C6F5)3 proceeded without any side reactions (cyclization, branching); however, only up to 60% monomer conversion was observed. PIPs with low

131 Mn (~1,200 g/mol) and Mw/Mn ~1.7 were obtained. NMR spectroscopy and MALDI-

Tof revealed that the unsaturation were almost exclusively in trans-1,4 configuration (92-

96.5%).

Figure 2.6.3 – Chemical scheme of cationic polymerization of IP with 131 1-(4-methoxyphenyl) ethanol as the initiator and B(C6F5)3 as the co-initiator.

This thesis conducts cross-disciplinary studies of NR biosynthesis. The

biochemistry of plants is distinctive from other genus of life because plant biochemical pathways tend to be particularly flexible and responsive to changes in the plant’s environment, both as a survival mechanism and as a mean of making optimum use of limiting resources.132 In the year 2000, enzymes are known to catalyze about 4,000 bio- transformations.133 We will investigate NR biosynthesis and bioemulative NR systems.

CHAPTER III

EXPERIMENTAL

3.1 Materials

Nerol (>98.0%, TCI America), geraniol (>96.0%, TCI America), 3,4- dihydropyran (DHP, 99%, ACROS Organics), pyridinium p-toluenesulfonate (PPTs,

>98.0%, TCI America), tert-butyl hydroperoxide (TBHP, t-BuO-OH, 70% in H2O, TCI

America), selenium dioxide (SeO2, 99.8%, Sigma Aldrich), sodium hydride (NaH, 95%,

Sigma Aldrich), were used as received. Triphenylphosphine (PPh3), tetrabromomethane

(CBr4), disodium dihydrogen pyrophosphate (Na2H2P2O7), ammonium hydroxide

(NH4OH), 40% (w/w) aqueous tetra-n-butylammonium hydroxide (NBu4OH), 1-bromo-

3-methyl-1-butene, tris(tetra-n-butylammonium) hydrogen pyrophosphate

((NBu4)3HP2O7, >97%), potassium tert-butoxide (95%), allyltrimethylsilane (ATMS,

98%), and 9-borabicyclo[3.3.1]nonane (9-BBN, 0.5 M in THF), hydrogen peroxide (30% w/w in H2O) were purchased from Aldrich and used as received. Salicylic acid, N-

bromosuccinimide (NBS, >99%), dimethyl sulfide ((CH3)2S, >99%) and p- toluenesulfonyl chloride (TsCl, >98.5%), divinyl adipate (DVA, > 98%) were purchased from TCI America and used as received. Celite was purchased from Fisher. Candida antarctica lipase B (CALB, immobilized on a macroporous acrylic resin, Novozym® 435,

Sigma) and Dowex AG 50W-X8 cation-exchange resin (100-200 mesh, hydrogen form) was purchased from Sigma. Tetrahydrofuran (THF, 98.5%), dichloromethane (CH2Cl2, 46

99%) and toluene (99.2%) were distilled over calcium hydride (Aldrich) before use wherever noted. Hexanes (98.5%) and ethyl acetate (≥ 99.5%) were purchased from

EMD and was distilled over sodium and benzophenone before use. Thin layer

chromatography (TLC) silica plates (Dynamic Absorbent Co.) and alumina plates

(Sigma-Aldrich) were used as received. 2,2-dimethyloxirane (IBEpx, >97.0%), 2-

methyloxirane (PPEpx, >99.0%) and di-t-butylpyridine (DtBP, >98.0%) were purchased

from TCI America and used as received. 2-methylpropene (isobutylene (IB), >99%) and

chloromethane (MeCl, >99.5%) were obtained from ExxonMobil. IB and MeCl were dried by passing the gas through a column filled with BaO and CaCl2. Titanium (IV)

tetrachloride (TiCl4, 99.9%) was purchased from ACROS Organics and from Sigma-

Aldrich and used as received. Methanol (99.8%) was purchased from Mallinckrodt

Chemicals and used as received. Sodium bicarbonate (99.7%) and anhydrous magnesium

sulfate (98.0%) were purchased from EMD Chemicals USA and used as received.

Tris-HCl (Tris(hydroxymethyl) aminomethane hydrochloride buffer was obtained

from Sigma-Aldrich and TCI America and used as received. Isoprene (IP, 99%),

stabilized with 100 ppm p-tert-butylcatechol, was obtained from Sigma-Aldrich and used

as received. HPLC grade tetrahydrofuran (THF) was obtained from Fisher Scientific and

was freshly distilled over sodium and benzophenone before use. HPLC grade THF was

also used for the SEC system.

3.1.1 Preparation of Washed Rubber Particles (WRP)

The latex of Hevea brasiliensis (RIMM 600 and IAC40) was collected from

research plantation trees at the Regional Centre of Votuporanga (São Paulo State, Brazil),

immediately stabilized, then shipped on dry ice to where the USDA (United States

Department of Agriculture) and stored at -80°C until use. WRP was prepared at the

USDA. The frozen latex was thawed to room temperature and placed on ice. Wash buffer

(100mM tris-HCl, 5 mM dithiothreitol (DTT) and 0.1 mM 4-(2-Aminoethyl)

benzenesulfonyl fluoride hydrochloride (AEBSF, pH = 7.5) was added to the latex and

the diluted latex was divided into 250 mL aliquots. The latex was then centrifuged at

3000 rpm for 10 min at 4oC. The top fraction was collected and transferred into fresh

wash buffer, and centrifuged at 5000 rpm for 10 min at 4oC. The same procedure was repeated with centrifugation at 7000 rpm for 10 min at 4oC. The top fraction was labeled

1X WRP. This washing/centrifugation procedure was repeated for the 1X WRP and the

sample was labeled as 2X WRP. Similar procedure was followed with 2X WRP to obtain

3X WRP. 3X WRPs were used for all in vitro experiments in this thesis. The WRPs were mixed with a small amount of wash buffer and 10% glycerol, and the latex was added into liquid nitrogen drop-wise using a pipette. The small beads forming in the liquid nitrogen were stored at -80°C to preserve the enzymatic activity. The latex and WRP samples were shipped to University of Akron on dry ice and stored at -80°C until use.

3.2 Procedures

Procedures for in vitro NR biosynthesis, model reactions, and preparation of the synthetic macroinitiator are discussed in the following sections.

3.2.1 in vitro Natural Rubber Biosynthesis (Hevea WRP) 3.2.1.1 Small Scale Synthesis (USDA)

In vitro experiments were carried out in wells of 96-well filter plates (Millipore

Durapore membrane 0.65 μm) using WRP-1. The reaction volume was 20 μL comprising of 2 μL of buffer (100 mM Tris-HCI, pH 7.5, 1.25 mM MgSO4, 5 mM DTT), 0.4 μL

(4×10-8 mol) of 100 mM IPP, 0.6 μL (6×10-8 mol) of 1mM FPP and 2 mg of WRP-1from

RRIM 600 in 17 μL of water. 5 hrs and 24 hrs reaction times were used at 25oC in an incubator. The reactions were stopped by adding 40 μL (3.2×10-6 mol) of 80 mM EDTA.

The filter plate was vacuumed and then washed two times with 150 μL water then once with 95% ethanol, then oven-dried at 37oC for 30 minutes.

3.2.1.2 Large Scale Synthesis (USDA)

One larger scale reaction was performed in 3.8 mL buffer (100 mM Tris-HCl, 1.25

-7 mM MgSO4, 5 mM DTT, 0.1 mM AEBSF, pH 7.5) containing 0.1 mM (3.8×10 mol) IPP,

15 μM (5.7×10-8 mol) FPP and 76 mg WRP-3 from RRIM 600. The reaction was run at room temperature with gentle stirring and was stopped by adding 40 mM EDTA (330 μl of 0.5 M

EDTA) after 24 hrs incubation. A blank was also prepared that had no FPP and IPP.

3.2.1.3. “Bioemulative” Experiments Using Synthetic Isoprene with WRP-3

(USDA)

Together with the reaction in 3.2.1.1 one reaction was performed in 3.8 mL buffer

(100 mM Tris-HCl, 1.25 mM MgSO4, 5 mM DTT, 0.1 mM AEBSF, pH 7.5) and 15 μM

(5.7×10-8 mol) FPP and 76 mg WRP-3 from RRIM 600, to which 26mg (0.38 mmol) IP was added. The reaction was run at room temperature with gentle stirring and was stopped by adding 40 mM EDTA (330 μl of 0.5 M EDTA) after 24 hrs incubation.

3.2.2 Experiments with IAC40 latex and WRP 3.2.2.1 Solids Content Determination for Latex and WRP

Solids content of IAC40 latex and WRP were determined by three different methods: 1) simply freeze drying the latex and WRP in vials without washing; 2) washing three times with buffer (100mM tris-HCl, pH = 7.5) and MQ water after self- coagulation, then freeze-dried to constant weight; and 3) by precipitating the NR latex and WRP into chilled methanol (4 oC). The precipitated polymer was white. The polymer was then washed with buffer (100mM tris-HCl, pH = 7.5) and MQ water to remove water-soluble solids and freeze-dried in the vacuum oven to constant weight.

3.2.2.2 In situ Raman monitoring (UAkron)

0.5 g active IAC40 WRP or IAC40 latex was sealed in glass vials (Fischer

Scientific, 1ml silanized glass vials) using a crimper. After the injection of isoprene (IP)

(0.102 g, 0.15 mL, 1.50 mmol) through the hermetic cap, the contents were vigorously shaken for one minute to ensure good mixing. The reactions were monitored by in situ

Raman spectroscopy. After incubation for 24 hours, the product was washed three times

with buffer (100mM tris-HCl, pH = 7.5) and de-ionized (DI) water. The rubber was then

freeze-dried until constant weight. The initial rubber weight was obtained by washing the

rubber, which was self-coagulated from latex/WRP at room temperature, three times with buffer (100mM tris-HCl, pH = 7.5) and de-ionized water, then freeze-drying till constant

weight. The initial rubber content data for the WRP and latex are based on the average of

three independent measurements (latex = 19.1% ± 0.4%, WRP = 17.8% ± 0.3%).

3.2.2.3 Micro-Raman Spectroscopy

The laser source was Lexel Raman Ion Krypton laser and the excitation wavelength was set at 647 nm. Raman signal was collected by a Horiba Jobin-Yvon

Labram HR single monochromator equipped with a nitrogen-cooled CCD camera. The sealed polymerization vial was fixed under a long-working distance 50X objective from

Mitutoyo with Numerical Aperture (NA) = 0.42. Spectra were collected for five minutes

for every data point, and then the laser was blocked for five minutes to avoid heating and fluorescence. This sequence was repeated for the duration of the first six hours of the experiment. The samples were incubated for a total of 24 hours. The position of the monochromator in most of the measurements was fixed at 1500 cm-1 to ensure the

capture of the major Raman modes characteristic of the C=C bonds in IP and PIP. The

schematic setup of the instrumentation is shown in Figure 3.2.1 and the actual

instrumentation is shown in Figure 3.2.2.

Figure 3.2.1 - Experimental set-up for the in situ Raman measurements.

a) b) Figure 3.2.2 -.Micro-Raman instrumentation in the Sokolov lab: a) before the experiment b) during in situ monitoring.

3.2.2.4 Experiments under CO2 atmosphere (UAkron)

1 mL silanized glass vials (Fischer Scientific), containing the latex or WRP??

Which ?, were sealed. For the experiments under CO2 atmosphere, the vials were purged with CO2 from dry ice and the pressure was regulated using an exit bubbler, as depicted in Figure 3.2.3. For samples denoted as KC_121109_W_IP1 (CO2/50X) and 52

KC_121409_W_IP3(CO2/50X) in Table 4.2.4, ~0.5 g of active IAC40 WRP or IAC40 latex and 0.10 g IP (0.15 mL, 1.50 mmol) were injected through the hermetic cap. For samples denoted as KC_120809_L_IP1 (CO2/50X) and KC_121109_W_IP2 (CO2/50X)

(Table 1), ~0.65 g of IAC40 WRP or IAC40 latex and 0.13 g of IP (0.19 mL, 1.95 mmol) were injected into the vials through the hermetic cap. The contents were then vigorously shaken for one minute to ensure good mixing. The rationale for increasing the reaction volume for the last two experiments (i.e. KC_120809_L_IP1 (CO2/50X) and

KC_121109_W_IP2 (CO2/50X)) was due to the horizontal configuration of the sample vials during Raman measurement. There were several incidences where the NR sample detached from the vial surface and the laser had to be re-aligned and re-focused when

placed in a horizontal position. By increasing the reaction volume, we hoped to prevent

detachment from the glass vial surface. It is important to note that the only variant was

the reaction volume. Therefore, the mole ratio between the latex/WRP and the IP

remained constant. After incubation for 24 hours, the products were washed three times

with buffer (100mM tris-HCl, pH = 7.5) and Milli-Q (de-

Figure 3.2.3 - Methodology to exchange the atmosphere within the closed vial.

3.2.2.5 “Bioemulative” experiments using deuterated isoprene (UAkron)

The deuterated isoprene(DIP) monomer (Oakridge National Lab, TN,

synthesized: 6/3/2008) was sealed in a vacuumed ampule and used as received. The

purity of the sample was determined by Gas Chromatography (GC) prior to use and the

sample was found to be 85.1% pure. 0.5 g active IAC40 latex were sealed in glass vials

(Fischer Scientific, 1ml silanized glass vials) using a crimper. 0.102g mixturesof isoprene

(IP) and D-IP (0.15 mL, 1.50 mmol) were injected into the vials through the hermetic cap with the following concentrations at 100% D-IP, 50/50 D-IP/IP and 100% IP. Two incubations were performed for each mixture. The contents were vigorously shaken for one minute to ensure good mixing. The 100% D-IP experiment was monitored by in situ

Raman spectroscopy. After incubation for 24 hours, the product was washed three times with buffer (100mM tris-HCl, pH = 7.5) and de-ionized water. The rubber was then freeze-dried until constant weight. The initial rubber weight was 19.1% ± 0.4% for latex and 17.8% ± 0.3% for WRP, which were determined previously as described in 3.2.2.1.

3.3 Synthesis of Macroinitiator

3.3.1 Synthesis of Protected Nerol (PN, product 2 in Figure 4.4.1 in Section 4.4)

A solution of nerol (10 g, 65 mmol) in dichloromethane (65 mL) was placed in a

250 mL one-neck round-bottom flask equipped with a condenser and a magnetic stirrer. di-hydropyran (6.55 g, 78 mmol) and pyridinium p-toluenesulfonate (1.63 g, 6.5 mmol) were added to the reactor and the resulting solution was stirred and thermostated at 27°C for 4 hours. Then the reaction mixture was concentrated using a rotary evaporator under reduced pressure, diluted with ethyl acetate and washed with saturated aqueous sodium 54

bicarbonate solution. The organic layer was dried over magnesium sulfate, filtered and

concentrated to yield transparent oil. The residue was purified by flash chromatography

(63-200 µm) eluent: ethyl acetate/hexane, 1:8 v/v; (silica TLC: Rf = 0.7) on silica gel to

yield PN (~14 g, 90%). The product was analyzed by 1H and 13C NMR spectroscopies.

3.3.2 Synthesis of Protected Nerol-OH (PN-OH, product 3)

A solution of PN (13 g, 55 mmol) in dichloromethane (70 mL) was placed in a

250 mL one-neck round-bottom flask equipped with a condenser and a magnetic stirrer.

tert-butyl hydroperoxide (24 mL, 173 mmol), selenium dioxide (0.61 g, 5.5 mmol) and

salicylic acid (0.76 g, 5.5 mmol) were added to the reactor and the resulting solution was

stirred at 30°C for 24 hours. Diethyl ether and water were added and the layers were

separated. The organic phase was washed with water three times. The combined organic

phase extracts were washed twice with saturated sodium bicarbonate aqueous solution,

dried over anhydrous magnesium sulfate, filtered, and concentrated by rotor evaporation.

The crude oil was purified by flash chromatography (63-200 µm, eluent: ethyl

acetate/hexane, 1:5 v/v; silica TLC: Rf = 0.15) on silica gel to yield PN-OH (~3 g, 25%)

(The literature reports ~30% yield for compounds with similar chemical structures103).

The product was analysed by 1H and 13C NMR spectroscopies.

3.3.3. Synthesis of Protected Nerol Tosylate (PN-Ts, product 4)

TsCl was recrystallized in ethyl acetate before use. In a 250mL round bottom flask, PN-OH (9.98g, 39mmol, a combination of pure PNOH from several syntheses),

BnNMe2 (0.522g, 3.86mmol), and K2CO3 (8.142g, 59mmol) were added in 30mL of

distilled water at room temperature. A stock solution of 1M KOH was made to maintain

pH of 10 during the reaction. The mixture was stirred vigorously and TsCl (11.250g,

59mmol) was added in 10 portions, every 6 minutes over the first hour. Immediately after

the addition of TsCl, drops of KOH were added to maintain the pH at 10. The pH was

monitored using a Mettler-Toledo portable digital pH meter. After the addition of TsCl,

the reaction was monitored with TLC (silica) and continued until there was no more PN-

OH (Rf = 0.1) spot. The Rf value of PN-Ts is 0.5 with 15:1 Hx/Ether as eluent. 1.00g of

N,N-dimethylethylene diamine was added to the mixture and stirred for 10 minutes to

react with the excess TsCl and quench the reaction. Water was added and the organic

phase was extracted with ethyl acetate. The organic fraction was washed with water and

brine two times and dried over anhydrous MgSO4. The crude product was purified by flash chromatography on silica gel to yield PN-OTs (12.62 g, 92%): silica TLC

(hexane/ether, 15:1, v/v) Rf = 0.5.

3.3.4 Synthesis of Polyisobutylene-Protected Nerol (PIB-PN, product 5)

In the glove box, sodium hydride (0.024 g, 1.0 mmol) and PIB-OH (5,800 g/mol,

Mw/Mn = 1.06, 1.02 g, 0.18 mmol) was added to 25 mL of distilled THF in a 100 mL

three-neck round-bottom flask. The mixture was stirred by a mechanical stirrer and PN-

Ts (1.2 mmol) was added drop-wise and the mixture was stirred for 5 days. Then the reaction mixture was slowly poured into water at 3°C, washed with a sodium sulfite solution (Na2SO3), and extracted 3 times with diethyl ether. The organic layers were washed twice with water, once with a saturated solution of sodium chloride, dried over anhydrous magnesium sulfate, filtered, and concentrated with a rotary evaporator. The polymer was analyzed by 1H and 13C NMR spectroscopies. The product was purified by

column chromatography (Hexane/Ethyl acetate, 10:1, v/v) three times due to tailing on

silica gel to yield PIB-PN (0.59 g, 60%).

3.3.5 Synthesis of Polyisobutylene-Nerol (PIB-Nerol, product 6)

A solution of PIB-PN (0.55 g, 0.09 mmol) in isopropyl alcohol (15 mL) and

hexane (7.5 mL) was placed in a 50 mL one-neck round-bottom flask equipped with a condenser and a magnetic stirrer and pyridinium p-toluenesulfonate (0.3 g, 1 mmol) was

added to the flask and stirred at room temperature for 20 hours. The polymer was precipitated in chilled methanol and washed with DI water three times. The polymer was freeze-dried. The residue was purified by column chromatography (eluent: Hx/EA, 10:1 v/v) on silica gel to yield PIB-Nerol (~0.53g, 98%). The product was analyzed by 1H and

13C NMR spectroscopies.

3.3.6 Synthesis of Polyisobutylene-Nerol-Bromide (PIB-Nerol-Br, product 7)

A solution of PIB-Nerol (0.50 g, 0.0DATF8 mmol) in dichloromethane (5 mL)

was prepared. In a 50 mL two-neck round-bottom flask equipped with a condenser and a

magnetic stirrer, dichloromethane (25 mL), NBS (0.888 g, 5.0 mmol) and Me2S (0.62g,

10 mmol) were added and chilled in a isopropyl alcohol/dry ice bath at -40 oC. A white

precipitate was formed at the bottom of the flask. The stirring speed was increased to

create a white suspension, and the solution of PIB-Nerol was injected through the rubber

septum. The reaction mixture was left stirring for one hour at –40 oC. The polymer was

precipitated in chilled methanol and washed with DI water and saturated sodium

bicarbonate solution three times. The polymer was then freeze-dried. The residue was

purified by column chromatography (eluent: Hexane/Ethyl aceate, 10:1 v/v) on silica gel

to yield PIB-Nerol-Br (~0.27g, 60%). The product was analyzed by 1H and 13C NMR spectroscopies.

3.3.7. Synthesis of Tris(tetra-n-butylammonium) Hydrogen Pyrophosphate

[(NBu4)3HP2O7]

The procedure was reported by Poulter et al.134 A solution of 3.33 g (15 mmol) of

disodium dihydrogen pyrophosphate Na2H2P2O7 in 41 mL of 0.104N aqueous solution of

ammonium hydroxide NH4OH (4.26 mmol) was passed through a column (Length:

20cm; Ø: 1cm) of Dowex AG 50W-X8 cation-exchange resin (100-200 mesh, hydrogen

form). The free acid was eluted with 90 mL of deionized water, and the resulting solution

(pH = 1.5) was immediately titrated to pH 7.3 with 40% (w/w) aqueous tetra-n-

butylammonium hydroxide NBu4OH. The resulting solution was concentrated under vacuum and then dried by lyophilisation to yield 13g of a hygroscopic white solid (96%).

The product was analyzed by 1H, 13C and 31P NMR spectroscopies.

3.3.8. Synthesis of Nerol-Pyrophosphate (Nerol-PP, model of product 8) This model reaction proceeded in the University of Bordeaux, France following

procedures adapted from Chen et al135. 0.50 g (3.3 mmol) of Nerol-Br and 3.48 g (4

mmol) of (Bu4N)3HP2O7 were solubilized in anhydrous acetonitrile under nitrogen atmophsere. The reaction was sitrred for 6 hours and then was concentrated under reduced pressure and a few drops of MQ water were added. The solution was then passed through Dowex AG50X8 ion exchange column (IXC) (NH4+ form). After the column the

solution was directly injected into LC (mobile phase: aceonitrile). Thedesired fration was

collected at the and and the product was freeze-dried into a white powder (0.94 g, 0.9

mmol, 27% conversion).

3.3.9. Synthesis of Protect Nerol-Divinyl Adipate (PN-DVA, Product 9)

Divinyl adipate (2.3 g, 12 mmol, [vinyl] 4 eq.) was added to an air-free, nitrogen-

purged, round-bottom flask containing PN-OH (Product 3, 1.01 g, 4.4 mmol), CALB (13

mg, 10% wt) and THF (1.5 mL). The mixture was stirred for 24 hours at 50 °C and

monitored by TLC (silica). After filtering off the enzyme with a 0.45 µm PTFE syringe

filter, the oil was concentrated with a rotary evaporator and purified by silica flash

chromatography twice. (elutent: Hexane/THF, 4:1, v/v, Rf = 0.5) The resulting oil had a

slightly yellow hue and the structure was analyzed by 1H and 13C NMR spectroscopies.

(Yield: 1.3g, 80%)

3.3.10 Synthesis of Trimethyl Pentyl Chloride (TMPCl, Product 10)

Sodium chloride (NaCl, 45 g, 0.77 mol) was placed into a 500 mL, three-necked, round-bottom flask. Concentrated sulfuric acid (65 mL, 0.65 mol) was added to the NaCl 59

very slowly from a dropping funnel to form HCl gas. The HCl gas was bubbled into neat

2,4,4-trimethyl-1-pentene (TMP-1, 11.6 g, 105 mol) in a round-bottom flask which was

kept in an ice-water bath. The reaction flask was connected to a trap and to another flask

containing concentrated sodium hydroxide solution to neutralize the unreacted HCl acid.

After 24 hours, TLC was checked and more NaCl and concentrated sulfuric acid was added to the setup until reaching 100% conversion observed by TLC over ~36 hours.

After work up, the reaction yielded 13.2 g of TMPCl (89 mmol, ~85%, loss of product from filtering)

3.3.11. Synthesis of Allyl Trimethyl Pentane (TMP-allyl, Product 11)

A 250-mL, three-necked, round-bottom flask equipped with an overhead mechanical stirrer was immersed in a hexane bath kept at -80 °C. TMPCl (4.01 g,

46mmol), DtBP (0.15 g, 2 mmol) were added to the reaction flask which was charged with a 60/40 (v/v) mixture of hexane (54 mL) and MeCl (36 mL). The reaction commenced with the rapid introduction of a pre-chilled stock solution of TiCl4 (10.1 g,

365 mmol). Amount of TiCl4 or ATMS used in this reaction was calculated depending on

the amount of TMPCl used (molar ratio [TiCl4]/[TMPCl] = 8). After stirring the reaction

mixture for 20 minutes, the intermediate was reacted with pre-chilled ATMS (6.21 g,

460mmol, molar ratio [ATMS]/[TMPCl] = 10). The solution was stirred for another 40

minutes in the glove box. The flask was taken out of the box and saturated aqueous

NaHCO3 solution was slowly added to the solution while the MeCl evaporated. The

solution was washed three times with distilled water and dried over anhydrous MgSO4.

The solvent was removed using a rotary evaporator and the oil was freeze-dried. (Yield:

4.12 g, conversion: 90%).

3.3.12. Synthesis of Trimethyl Pentane-OH (TMP-OH, Product 12)

Hydroboration followed by alkaline oxidation is the general method used to

convert PIBs with allyl (PIB-CH2-CH=CH2) end groups to the corresponding PIBs with

primary alcohol functionalized end groups. The same procedure was applied for TMP-

allyl. TMP-allyl (4.00 g, 24 mmol) was dissolved in distilled THF (100 mL) and added

drop-wise to an ice chilled THF solution of 9-BBN (17.8 mL, 8.9 mmol, 0.5 mol/L)

under an inert atmosphere (N2) over 30 minutes. After stirring the reaction mixture at

room temperature for five hours, 25% w/w KOH solution in methanol (1.5 g in 6 mL of

methanol) followed by 30% v/v H2O2 solution(2.7 mL) were added drop-wise to the

reaction mixture while maintaining the reaction temperature at 3 ºC. The reaction

mixture was then allowed to react for 2 hours. Hexane (100 mL) was added to the

solution, stirred for five minutes and poured into 200 mL saturated potassium carbonate

solution. The organic layer was washed several times with distilled water and dried with anhydrous MgSO4. The solution was filtered and rotary evaporated to dry. (Yield: 3.12 g,

conversion: 72%).

3.3.13. Synthesis of Protected Nerol-Divinyl Adipate-Trimethyl Pentane (PN-DVA-TMP, Product 13)

TMP-OH (0.78 g, 4.6 mmol) was added to an air-free, nitrogen-purged, round-

bottom flask containing PN-DVA (2.03 g, 5.0 mmol), CALB (17 mg, 10% wt) and THF

(10.0 mL). The mixture was stirred for 24 hours at 50 °C and monitored by TLC. After

filtering off the enzyme with a 0.45 µm PTFE syringe filter, the oil was concentrated with

a rotary evaporator and purified by silica flash chromatography twice. (elutent:

Hexane/THF, 4:1, v/v, Rf = 0.5) The chemical structure of the transparent resulting oil was analyzed by 1H and 13C NMR spectroscopies. (Yield: 1.84 g, 3.45 mmol, 70%)

3.3.14. Synthesis of Nerol-Divinyl Adipate-Trimethyl Pentane (Nerol-DVA-TMP,

Product 14)

A solution of PN-DVA-TMP (1.05 g, 2.0 mmol) in isopropyl alcohol (10 mL) and

hexane (5 mL) was added to a 25 mL one-neck round-bottom flask equipped with a

condenser and a magnetic stirrer. Pyridinium p-toluenesulfonate (PPTs, 0.2 g, 0.8 mmol)

was added to the reaction mixture and the reaction was stirred at room temperature for 20

hours. The reaction mixture was concentrated with a rotary evaporator which led to the

formation of white crystals of pyridinium p-toluenesulfonate at the bottom of the flask.

Petroleum ether was added to dissolve the product but not the PPTs. The product was

then washed with DI water three times. The product was purified by silica flash

chromatography (eluent: Hexane/THF, 4:1, v/v) on silica gel to yield N-DVA-TMP

(Yield = 0.8 g, 90%). The product was analyzed by 1H and 13C NMR spectroscopies.

3.3.15. Synthesis of Nerol-Bromide-Divinyl Adipate-Trimethyl Pentane (Nerol-Br-DVA

-TMP, Product 15)

The bromination of Nerol-DVA-TMP to Nerol-Br-DVA-TMP proceeded in

dichloromethane at -40 oC using N-bromosuccinimide (NBS) and dimethyl sulfide.

Nerol-DVA-TMP (0.55 g, 1.2 mmol) and NBS (0.27 g, 1.5 mmol) were added to CH2Cl2 and left stirring for 20 minutes before warming the reaction flask to 0 oC. After the

o temperature stabilized at 0 C, dimethyl sulfide (Me2S, 0.1 g, 1.6 mmol) was injected and

the reaction proceeded for 6 hours. The reaction yielded ~0.4 g (0.8 mmol) of Nerol-Br-

DVA-TMP, which corresponded to ~65% conversion. The product was purified by

column chromatography (eluent: Hexane/Ethyl acetate (EA), 10:1, v/v) (Rf = 0.25 for

Nerol-Br-DVA-TMP, Rf = 0.1 for Nerol-DVA-TMP).

3.3.16. Synthesis of Nerol-Pyrophosphate-Divinyl Adipate-Trimethyl Pentane (Nerol-PP-

DVA-TMP, Product 16)

The phosphorylation of Nerol-Br-DVA-TMP to Nerol-OPP-DVA-TMP proceeded in air-free conditions for 6 hours, which was in accordance with the literature.

Nerol-Br-DVA-TMP (0.3 g, 0.6 mmol) and (Bu4N)3HP2O7 (0.9 g, 1 mmol) was added to

2 mL of anhydrous acetonitrile equipped with a magnetic stirring bar. After 6 hours, the anhydrous acetonitrile was evaporated under reduced pressure and a few drops of MQ water were added. The solution was then passed through Dowex AG50X8 ion exchange column (IXC) (NH4+ form).

3.3.17. Synthesis of Nerol-P(O)(OEt)2 (Model Reaction for Phosphorylation).

A solution of Nerol (250 mg, 1.62 mmol) and pyridine (0.16g, 2.02 mmol) in chloroform (5 mL) was added to a 10 mL one-neck round-bottom flask equipped with a

15 mL additional funnel and a magnetic stirrer. The reaction mixture was chilled to 1 oC

using an ice bath. Diethyl chlorophosphate solution (0.28 g, 1.62 mmol in 1 mL CHCl3) was added to the flask slowly using the additional funnel and the reaction mixture solution was stirred at 1 oC for 4 hours. The reaction was monitored by TLC (silica) (1:3

Hexane/Ethyl acetate). The reaction mixture was concentrated slightly using rotary

evaporator and then loaded directly onto a silica gel column (1:3 Hexane/Ethyl acetate) to

separate and purify Nerol-P(O)(OEt)2 (~150 mg, 32%, Rf = 0.34) and Nerol (~125 mg,

50%, Rf = 0.9). Nerol-P(O)(OEt)2was a light yellow oil with some crystals. The product

was analyzed by 31P NMR spectroscopies.

3.3.18. Synthesis of Nerol-P(O)(OEt)2-DVA-TMP (Product 15).

A solution of Nerol-DVA-TMP (0.2 g, 0.44 mmol) and pyridine (0.05g, 0.65

mmol) in chloroform (3 mL) was added to a 5 mL one-neck round-bottom flask equipped

with a 15 mL additional funnel and a magnetic stirrer. The reaction mixture was chilled

to 1 oC using an ice bath. Diethyl chlorophosphate solution (0.1 g, 0.69 mmol in 1 mL

CHCl3) was added to the flask slowly using the additional funnel and the reaction mixture solution was stirred at 1 oC for 4 hours. The reaction was monitored by TLC (silica) (1:3

Hexane/Ethyl acetate). The reaction mixture was concentrated slightly using rotary

evaporator and then loaded directly onto a silica gel column (dichloromethane) to separate and purify Nerol-P(O)(OEt)2-DVA-TMP (~150 mg, 32%, Rf = 0.6) and Nerol

(~115 mg, 50%, Rf = 0.25).

3.3.19. Synthesis of Nerol-PP-DVA-TMP (Product 16).

Under inert atmosphere, a solution of Nerol-P(O)(OEt)2-DVA-TMP (0.15 g, 0.35

mmol) (Bu4N)3HP2O7 (0.8 g, 0.7 mmol) in chloroform (3 mL) was added to a 5 mL one- neck round-bottom flask equipped with a 15 mL additional funnel and a magnetic stirrer.

The reaction was stirred at room temperature for 2 days and was monitored by TLC

(silica) (acetonitrile). The reaction mixture was concentrated slightly vacuum then loaded directly onto a cellulose Whatman CF-11 column (anhydrous acetonitrile) to separate and

purify Nerol-P(O)(OEt)2-DVA-TMP (Rf = 0.4) and Nerol-PP-DVA-TMP (~240 mg,

54%, Rf = 0.7).

3.4. Laboratory Techniques and Instrumentation 3.4.1. Air-free Technique

The removal of contaminants (e.g. O2, H2O, CO2) and volatile impurities from the

glassware, solvents, and reagents used was necessary to avoid unwanted reactions for

some steps. This was accomplished by using a Schlenk line and air-free glassware. The

Schlenk line consisted of a 4-port air-free vacuum manifold made of Pyrex® glass tubing.

All connection ports used were Teflon stopcocks. A vacuum pump was connected to one

end of the manifold. A liquid nitrogen trap was placed between the manifold and the

pump to protect the pump oil from solvent contamination. Nitrogen gas (Praxair,

technical grade) was introduced into the vacuum line through the other end of the

manifold. The pressure of the nitrogen gas in the vacuum line was controlled using a

bubbler. A glass drying column packed with anhydrous calcium sulfate and dried

molecular sieves (4Å) was placed between the gas cylinder and the manifold to dry the

nitrogen gas.

3.4.2. Thin Layer Chromatography (TLC)

Silica plates (Dynamic Absorbent Co.) or alumina plates (Sigma-Aldrich) were

used to perform TLCs using different eluents depending on the compounds involved.

Specific conditions are given in the syntheses section. The typical solvents used were

hexanes, ethyl acetate, THF, and diethyl ether. Capillaries were made from disposable

pipettes over propane torch. For a typical TLC experiment, a drop of sample solution was

placed on the TLC plate using a capillary and the plate was eluted with the appropriate

solvent(s). The spots were observed using any of the following three methods : (a)

developing the plate by dipping in a solution of phosphomolybdic acid in ethanol

(Aldrich) (b) exposing it to a UV lamp (λ=254 nm) or (c) using an iodine chamber.

3.4.3. Column Chromatography

Column chromatography was used for the purification of compounds. The columns were packed with silica gel as follows: the appropriate eluent (specific conditions are given in synthesis details of each compound) was added to a beaker containing silica gel (Dynamic Adsorbent Inc, 63-200µm, silica gel classic column (Lot#

LB1708)) or alumina (Dynamic Adsorbent Inc, 63-200µm Alumina for DCC). The resulting slurry was quickly poured into a Pyrex® column and the solvent was allowed to drain through the column until its level was just above the surface of silica gel. The column was gently tapped during this time in order to insure that there were no air bubbles in the packed silica gel or alumina. After the column was uniformly packed, the crude product dissolved in a small amount of solvent was carefully loaded onto the column using a pipette. The solution was allowed to drain until the level reached the top of silica gel. A small amount of sand was added carefully to protect the top of the silica column. The eluent was run through the column applying air pressure at the top of the column, resulting in rapid separation of the products and high column performance. In flash chromatography, air pressure was applied. The fractions were analyzed by TLC and the fractions containing the pure product were combined and concentrated using a rotary evaporator under reduced pressure.

3.4.4 Size Exclusion Chromatography

Molecular weights (MWs) and molecular weight distributions (MWDs) were

determined by SEC. The SEC system used in this study was a Waters setup equipped

with six Styragel columns (HR0.5, HR1, HR3, HR4, HR5, and HR6) thermostatted at

o 35 C as the stationary phase. Tetrahydrofuran (THF) continuously distilled from CaH2 was used as the mobile phase at a flow rate of 1 mL/min. The list of detectors include a

Wyatt Technology Viscostar viscometer (VIS), a Wyatt Optilab DSP refractive index

(RI) detector thermostatted at 40oC, a Wyatt DAWN EOS 18 angle multi-angle laser light

scattering (MALLS) detector, a Wyatt quasi-elastic light scattering (QELS) and a Waters

2487 Dual Absorbance ultraviolet (UV) detector. (Note: the UV detector was used;

however, the signal intensity was very low to be useful as the injection concentration was

very low. This issue will be addressed in future) Absolute molecular weights and radii of

gyration were determined using ASTRA® V software 5.3.4 and dn/dc = 0.130 reported

for high cis-poly-isoprene.8 The dn/dc value was confirmed with polyisoprene standards

(Mn = 2,450 and 9,870 g/mol and Mw/Mn = 1.02 and 1.03) supplied by Scientific Polymer

Products, Inc. The schematic of the SEC system is shown in Figure 3.4.1. In our experiments, it is important to note that the viscometer was not used because the protein residues left in the rubbers clog the capillaries of the viscometer.

Figure 3.4.1. High resolution SEC system at the Puskas Lab.

3.4.5. NMR sample preparation

The dried rubber samples incubated with IP were dialyzed using Nest Group’

SpinDIALYZER (50 μL) with 0.6 μm PTFE membranes (Millipore) in order to separate the insoluble gel fraction and the soluble component. The solution outside the dialysis chamber and the gel within the chamber were concentrated and freeze-dried until constant weight. After freeze-drying, the soluble fraction from the outside chamber (~50-

70 mg) was dissolved in 1 mL of benzene-D6 (Cambridge Isotopes Inc.) or 1 mL of

toluene-D8 (Cambridge Isotopes Inc.) in a 3 mL glass vial. The samples were left to

dissolve in the dark for 6 hours and transferred to NMR tubes.

After incubation with D-IP and freeze drying, ~100 mg of the NR was dissolved

in 4 mL benzene-D6 and then filtered into a pre-weighed vial. The sample was dried with

a vacuum pump (Yield: ~50 to 60 mg after drying) and ~1 mL benzene-D6 was added to

make a ~50 mg/mL solution.

3.4.6. 1H NMR procedure

1H NMR was performed using either a 300MHz Varian Mercury NMR or a 500

MHz Varian One NMR instrument 128 scans were taken with d1 (relaxation time) = 10

seconds and 45° pulse angle. The temperature was controlled at 25 °C and the solvents

used were typically benzene-D6, toluene-D8 or chloroform-D. Concentrations of the

sample solutions used for the 1H NMR measurements are listed by the figures. Total

acquisition times are listed in the appendix for each NMR spectrum.

3.4.7. 13C NMR procedure

13C NMR was performed using either a300MHz Varian Mercury NMR or a 500

MHz Varian One NMR instrument. 10,000 scans were taken with d1 (relaxation time) =

10 seconds and 90° pulse angle, as suggested by Tanaka et al.136 The temperature was

° controlled at 25 C and the solvents used were typically benzene-D6, toluene-D8 or

chloroform-D. Concentrations of the sample solution used for the 13C NMR

measurement are listed by the figures.

3.4.8. Gas Chromatography

Gas chromatography (Shimadzu GC-8A) was performed using an Equity-1 fused

silica capillary column, a TCD detector, and a CR501 recorder using He as a carrier gas.

The temperature range was between 80°C to 280 °C and the inject-to-collection delay was

1 minute.

3.4.9. Matrix Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry

(MALDI-ToF MS)

MALDI-ToF mass spectra were acquired on a Bruker UltraFlex-III ToF/ToF mass

spectrometer (Bruker Daltonics, Inc., Billerica, MA) equipped with a Nd:YAG laser (355

nm) for verifying the purity and mass distribution of the products. All spectra were measured in positive reflector mode. The instrument was calibrated prior to each measurement with an external poly(methyl methacrylate) (PMMA) standard. Individual solutions of polymer (10 mg/mL) in anhydrous THF (99.5 %, Aldrich), 1,8,9- trihydroxyanthracene matrix (dithranol, 20 mg/mL, 97 %, Alfa Aesar), and silver trifluoroacetate (AgTFA, 10 mg/mL, 98%, Aldrich) in anhydrous THF were mixed in the ratio of polymer:matrix:cationizing salt (1:10:2), and 0.5 µL of the resulting mixture were deposited on microtiter plate wells (MTP 384-well ground steel plate). After evaporation of the solvent, the plate was inserted into the MALDI source. The spectra were obtained in the reflectron mode at an acceleration voltage of 20 kV. The attenuation of the

Nd:YAG laser was adjusted to minimize unwanted polymer fragmentation and to maximize the sensitivity.

3.4.10. Electrospray Ionization Mass Spectrometry (ESI-MS)

ESI mass spectra were acquired with a Bruker Daltonics Esquire-LC ion trap mass spectrometer for the identification of mass. The sample was dissolved in anhydrous

THF (99.5 %, Aldrich) at 1µg/µL, and the resulting solutions were mixed with 1µg/µL solution of sodium trifluoroacetate (98%, Aldrich) as a cationizing agent in THF in the ratio 100:1(sample:salt) (v/v). Experimental conditions: positive mode; dry gas: nitrogen 70

(8 L/min); drying temperature: 300 °C; nebulizer gas: nitrogen (10 psi). The sodium salt solutions were introduced into the ESI source by direct infusion using a syringe pump at a flow rate of 250 µL/h.

CHAPTER IV

RESULTS AND DISCUSSION

4.1. In vitro Natural Rubber Biosynthesis

4.1.1. Monitoring the Growth of in vitro Natural Rubber by High-Resolution Size

Exclusion Chromatography (HR-SEC)

Enzymatically active WRP (WRP-1) was isolated from RRIM600 latex as

described in Chapter 3.1.2 RRIM600 stands for Rubber Research Institute of Malaysia –

600, which is a representation of a typical H. brasiliensis clone. The solids content was

measured by drying the WRP-1 at 60 oC for 24 hours, and was found to be 278 mg/mL

(27.8 wt%). The enzymatic activity was determined by procedures in Chapter 3.1.2, in

which the WRP was incubated with 14C-labeled IPP with and without FPP initiator.

Figure 4.1.1 shows the well plates used for in vitro NR biosynthesis. Mau and Cornish112 developed multi-well plate assay to determine the enzymatic activity rapidly. Previously, enzymatic activities of prenyltransferases were determined by incubating the rubber particles with radiolabelled IPP in each individual micro-centrifuge tubes with varying concentration of IPP at a saturated concentration of FPP (or APP initiator).113

Figure 4.1.1 – Micro well-plates in which in vitro NR biosyntheses are performed.

In the multi-well method, PVDF filters were placed at the bottom of the wells. In each micro-well, 2 mg WRP samples were incubated with FPP initiator, 14C IPP monomer, and Mg2+ cofactor. (Figure 4.1.2a) After 4 hours, EDTA was added to terminate the reactions. EDTA is a ligand that strongly chelate with metal co-factors

(Mg2+) to quench the reaction. The buffer in the wells was suctioned out of the wells by vacuum and washed repeatedly with water, 1M HCl and 95% ethanol to remove unreacted 14C labeled IPP. (Figure 4.1.2b) The filters were taken out of the wells and placed in scintillation fluid for radioactivity measurements to determine the enzymatic activity.

Figure 4.1.2 – An example micro-well and its constituents a) during incubation, b) after incubation.

Using this method, it was found that RRIM600 WRP-1 was able to incorporate

10.3 µmol 14C-labeled IPP per g of dry rubber when incubated with non-labeled FPP synthetic initiator (Figure 4.1.3). In the absence of FPP, 3.0 µmol 14C -labeled IPP per g of dry rubber incorporated. These data agree with activities reported in the literature.118

Figure 4.1.3 – Enzyme activity measurement of RRIM600 WRP-1 using 14C IPP (USDA).

74 Figure 4.1.4 shows the SEC Refractive Index (RI) trace of the toluene-soluble

fraction of WRP-1 from RRIM600. The high molecular weight (MW) region shows a

peak in the 106 g/mol range (1H) with a shoulder at ~105 g/mol. There are four distinct

low MW peaks (i.e. labeled as 1L, 2L, 3L, and 4L) in the spectrum. The MW of these

peaks approximately corresponds to 2, 3 and 4 times that of the 1L peak. Since the RI

signal is proportional to concentration, by integrating the area under the RI traces, the

ratio of high MW rubber and low MW constituents can be obtained. The low MW

fraction was found to constitute ~50% of the toluene-soluble fraction. Table 4.1.1 lists the

MW data. Peak 1H is the average MW data given from the MALLS detector while peaks

1L~4L are peak MW data calculated from a MW-elution time calibration curve

extrapolated from log MW-log elution time graph. Low molecular weight (4,000 Da) NR

was observed earlier by Archer et al. and Asawatreatanakul et al.3,8 Our high resolution

SEC analysis is of particular interest in our pursuit to understand natural rubber biosynthesis because of the identification of a high fraction of low MW intermediates.

Figure 4.1.4 – SEC RI trace of WRP-1.

75 Table 4.1.1 – HR-SEC data of WRP-1. a a b b b b 1H Mw/Mn 1H 1L 2L 3L 4L

(g/mol) Rg (nm) (g/mol) (g/mol) (g/mol) (g/mol)

WRP-1 1.63x106 2.12 80.5 500 900 1,600 2,400 a determined from MALLS detector. b determined from MW-elution time calibration curve.

Figure 4.1.5 presents the conformation plot (log Rg vs log Mn) for the high MW region of the toluene-soluble fraction isolated from WRP-1. From the plot, the slope of

0.539 suggests that this is a linear polymer. NR is known to have long-chain branched molecules. Field-Flow Fractionation (FFF) Chromatography of NR with 50% gel content showed long chain branching/gel (slope of 0.3~0.4).5

Figure 4.1.5 – Conformation Plot of WRP-1.

With enzymatically active WRP-1 from RRIM600, several in vitro experiments were conducted at the USDA under conditions that mimic the in vivo environment of the

76 cytosol. More specifically, the FPP initiator and the IPP monomer were incubated with

WRP-1, which contained the membrane-bound cis-prenyl transferase enzyme in buffer solution. The in vitro reactions were terminated after 5 and 24 hours by the addition of

Ethylene-Diamine-Tetraacetic-Acid (EDTA). EDTA is a ligand that strongly chelates with metal co-factors (Mg2+ or Mn2+). The presence of free co-factors to coordinate with the pyrophosphate end-groups is necessary for the activation of the dormant chain-end

and the polymerization.

Figure 4.1.6 compares the RI traces of representative 5 and 24 hour-incubated

samples with WRP-1. The 1H peak of WRP-1/24 shifted to higher molecular weight

relative to WRP-1 and WRP-1/5, and a distinctive second peak (2H) appeared with an

approximate MW of ~5×105 g/mol (Figure 4.1.6b). This observation is similar to the SEC traces reported in the literature when radio-labeled 14C-IPP monomer3 was used. For this

particular experiment, the weight fraction of the high MW materials (from integration of

the RI signals) increased to ~80% from 50% in the WRP-1. The evidence of MW growth

is supported by the increase of Rg and the increase of the high MW mass fraction. The Rg increased from 80.5 nm of the WRP-1 to 93 nm in WRP-1/24. Therefore, the 2H peak was identified to be newly-formed NR.

77 b) a) 1.2 1H WRP-1 1.0 WRP-1 WRP-1/5 1H 1.0 WRP-1/5 WRP-1/24 0.8 WRP-1/24 0.8

0.6 0.6 2H 2H 0.4

Relative scale 0.4 Relative scale 0.2 0.2

0.0 0.0

5 6 7 105 106 107 10 10 10 Molecular Weight (g/mol) Molecular Weight (g/mol)

Figure 4.1.6 – SEC of WRP-1, WRP-1/5 and WRP-1/24. High MW region. a) LS trace, b) RI trace.

The low MW fractions of the in vitro rubbers also show interesting changes.

Peaks 1L, 2L and 3L diminished with the rise of a new low MW peak, 5L, shown in

Figure 4.1.7. The peak MWs for the well-resolved low fractions are shown in Table 4.1.2.

1.2 WRP-1 2L WRP-1/24 1.0

0.8

0.6 4L 5L Relative scale 0.4 1L 3L 0.2

0.0 103 104 Molecular Weight (g/mol)

Figure 4.1.7 – SEC RI trace of WRP-1 and WRP-1/24. Low MW region. 78 The MW of 2L is ~900 g/mol, while peaks 4L and 5L correspond to 2,200 and 3,500

g/mol.

Table 4.1.2 – Approximate MW of peaks 1L to 5L from HR-SEC. 1L 2L 3L 4L 5L Sample ID (g/mol) (g/mol) (g/mol) (g/mol) (g/mol)

WRP-1 500 900 1,600 2,400 -

WRP-1/5 400 900 - 2,100 3,300

WRP-1/24 400 900 - 2,100 3,200 b determined from MW-elution time calibration curve.

NR latex from Hevea (H600 clone) was fractionated and the low MW fractions (<

4000 Da) were analyzed by MALDI-ToF MS. Figure 4.1.8 presents the MALDI-ToF

mass spectrum. The presence of AgTFA salt led to the formation of singly charged ions

(Ag+ charge) in the rage of 500-3000 Da. (approximately materials from L4 and L5 from

the HR-SEC.) The mass difference between the peaks was found to be 68 Da,

corresponding to the molecular weight of one polyisoprene repeat unit. From Figure

4.1.8, two distributions of oligoisoprenes can be observed.

Figure 4.1.8 – MALDI-ToF spectrum of fractionated low MW Hevea NR. (H600 clone).

Based on the m/z values of the [M+Ag]+ ions observed, the major distribution has

end groups of 32 Da, whereas the minor distribution has end groups of 88 Da. The

identification of the structure of these end groups is in progress. Possible end groups with

a mass of 32 Da are CH3OH. Possible end groups with a mass of 88 Da are

(CH3)2CHCH2CH2OH.

In addition to monitoring the growth of NR by HR-SEC, we also wanted to

perform gravimetric analysis. In order to do that, we needed to produce in vitro NR at a

‘larger’ scale. 76 mg WRP-3 from RRIM600 (isolated on a different day than WRP-1)

was incubated with synthetic IPP and FPP in a small centrifuge tube for gravimetric

80 analysis. After 24 hours the final material was freeze-dried to constant weight. The blank was also dried. Table 4.1.3 shows the gravimetric results. Under the conditions used

(shown under Table 4.1.3), substantial mass gain was observed in the sample incubated with IPP and FPP, relative to the blank. The mass gain is more than the IP equivalent of the IPP. However, it should be noted that the experimental error can be high because of the small scale.

Table 4.1.3 – Gravimetric analysis of WRP-3/24. Initial WRP Final Rubber Mass Gain Sample ID (mg) (mg) (mg) KC_030209_WRP (Blank) 76 52 WRP-3/24 104 52 3.8 mL of buffer, 76 mg of WRP + 112 mg IPP (32 mg IP equivalent) + 22 mg FPP (6 mg IP equivalent)

After 24 hours incubation with FPP and IPP, the gel content measured by dialysis reduced slightly from 33wt% to 28wt% (Table 4.1.4). This finding also supported the presence of newly formed soluble in vitro NR.

Table 4.1.4 – Gel fraction analysis of WRP-3 and WRP-3/24. Gel fraction Sample ID (%) WRP-3 33 WRP-3/24 28

The SEC traces and data are presented in Figure 4.1.9 and Table 4.1.5, respectively. In the high MW region, a slight shift of the peak towards higher MW can be seen, but 2H was not observed in this experiment. The Rg after incubation increased and

81 the relative amount of the high MW fraction decreased. Similarly to WRP-1 (Figure

4.1.5), the conformation plot had a slope of 0.54, characteristic of linear polymers.

a) b) WRP-3 0.30 1.2 WRP-3/24 WRP-3 WRP-3/24 4L 0.25 1.0 1H 0.20 0.8 0.15 0.6

Relative scale 0.10

Relative scale 0.4 2L 0.05 0.2

0.0 0.00 5 6 7 103 104 10 10 10 Molecular Weight (g/mol) Molecular Weight (g/mol)

Figure 4.1.9 – SEC traces of WRP-3 before and after incubation. (a) RI traces of low MW region, (b) RI traces of high MW region.

Table 4.1.5 - SEC data of WRP-3 and WRP-3/24.

1H Peak time 1H Peak MW Rg % Mass of Sample ID 6 High MW (min) (10 g/mol) (nm)

WRP-3 35.09 1.28 90 50 WRP-3/24 34.85 1.71 97 40 The retention time difference relative to the WRP-1 set is due to a system pump upgrade performed between the analyses.

In the low molecular weight region, the disappearance of the lowest MW peak,

2L, at around 1,400 g/mol can be observed. Although our SEC analysis showed that the

low MW regions vary within different batches of WRPs made from the same RRIM600 latex, 2L always decreased and 4L increased in the in vitro experiments for all incubations using WRP-1 and WRP-3. (The individual SEC traces are in Appendix B) It

82 had previously been shown that in-vitro NR biosynthesis was able to extend pre-existing

short-chain NR.8

In summary, NRs synthesized in vitro were analyzed by high resolution SEC. The

soluble rubber had approximately 50% high MW fraction between 105 and 3x106 g/mol, and 50% low molecular weight components with MW between 400 and 4,000 g/mol. In the presence of both FPP initiator and IPP monomer, high-resolution SEC was able to resolve the growth of both high and low MW rubber. Further, the formation of new NR was also indicated by mass balance measurements, which showed a net mass gain.

4.2. Substitution of the Isopentenyl Pyrophosphate (IPP) Monomer with Synthetic

Isoprene (IP)

The IPP monomer is produced from carbohydrates in rubber-producing plants as it was discussed in the Introduction. IP is also produced actively by plants and animals.

When excess IPP is present, the equilibrium shifts toward DMAPP which in turn is converted into IP by the isoprene synthase enzyme. IP is an ideal compound to remove excess IPP via the shifts in the equilibria (see Figure 2.3.4) for its ease of evaporation

(boiling point of IP = 34 oC).137 In addition, it is postulated that IP provides heat

protection for plants.138 Efforts in biological research are actively studying the

fundamental processes involved in the bacterial breakdown of terpenoids.139 Since many of the enzymatic reactions are reversible140, Puskas et al. hypothesized that IPP could be

generated in situ by “flooding” the active NR producing latex with IP, and a US patent

83 application has been filed in this regard. The following experiments were carried out to

investigate this hypothesis in detail.

4.2.1. In vitro NR Biosynthesis in the Presence of Synthetic IP

The WRP-3 described in Chapter 4.1 was incubated with FPP as initiator and IP as monomer. No IPP was present in the incubation experiments. The experimental

conditions are listed in Table 4.2.1 below and gravimetric analysis was performed. Since

IP is volatile and the only reagent added to the system, the mass gained after incubation was considered to arise from IP being converted into solids, presumably PIP.

Table 4.2.1 - Experimental conditions for WRP-3 with IP. Sample FPP IPP IP Rxn time (hr) KC_030209_ + - + 24 WRP-3/24(IP) (15μM) (100mM) *3.8 mL of buffer, 76 mg of WRP + 26 mg IP + 22 mg FPP 100 mmol 1L 68.12 g Wt. of IP =   3.8mL   25.9mg L 1000 mL mol

In this experiment, the weight of the IP incubated was calculated as 26 mg as seen

in the footnote of Table 4.2.1. Therefore, the mass gain of in vitro NR biosynthesis can be

used to approximate the conversion of IP into PIP. The weight of the rubber reported

was determined by the mass obtained after drying the samples in the vacuum oven over

five days till constant weight.

84 Table 4.2.2 – Gravimetric analysis summary for experiments with WRP-3. Sample Initial Rubber Wt. diff Conversion WRP (mg) Wt. (mg) (mg) (%) KC_030209_WRP (Blank) 76 52 KC_030209_WRP-3/24(IPP) 52 104 +52 88 KC_030209_WRP-3/24(IP) 52 103 +51 100 (196)

From Table 4.2.2, it was observed that there was a positive mass difference between initial and final polymer weight; suggesting a polymerization reaction has taken place. Further, it was observed that KC_030209_WRP-3/24(IPP) had similar mass gain

compared to KC_030209_WRP-3/24(IP), which was incubated with IP as the monomer.

After the observation that WRP-3 isolated from RRIM600 incubated with

synthetic IP in an in vitro NR biosynthesis showed mass gain, the next set of

experiments involved WRP and raw latex from IAC40 Hevea. IAC40 stands for Instituto

Agronômico de Campinas clone #40 from São Paulo State, Brazil. This particular

genotype carries high NR yield genes. The reason behind switching the genotype was that

our collaborators at the USDA did not have enough stock of RRIM600 Hevea. The

enzymatic activity of IAC40 WRP was determined by procedures described in Chapter

3.1.2, in which the WRP was incubated with 14C-labeled IPP with and without FPP

initiator. The resulting rubber was then isolated and subject to scintillation spectroscopy

to determine the enzymatic activity by the amount of radioactivity. It is important to add

additional initiator when determining enzymatic activity because it was found that the

omission of initiator typically inhibited the enzymatic activity.112 FPP is chosen as the

initiator in this case because it is one of the most effective initiators for Hevea WRPs116 and it is typical to observe a multiple fold increase in enzymatic activity when FPP is added in the assay. Our collaborators at the USDA used the procedures established by

85 Cornish et al.112 and determined that IAC40 WRP was able to incorporate 12.7 µmol/g of

14C -labeled IPP when incubated with FPP synthetic initiator (Figure 4.2.1). When compared to RRIM600 WRP, which could incorporate 10.3 µmol/g dry rubber / 4 hour at

25oC under the same procedure, IAC40 WRP was found to be slightly more reactive.

Figure 4.2.1 – Enzyme activity measurement using 14C-labelled IPP (USDA).

In order to analyze the samples by gravimetric analysis, the solids contents of the starting latex and WRP samples were measured by three different methods. Table 4.2.1a and b show solid content data determined by freeze drying the latex and WRP in vials without washing. Six samples, three latexes and three WRPs were dried to constant weight.

Table 4.2.3a and b show the data for 6 samples (3 IAC40 latex and 3 WRPs) washed three times with buffer (100mM tris-HCl, pH = 7.5) and DI water (distilled, deionized water) after self-coagulation, then freeze-dried to constant weight.

86 Comparison of Tables 4.2.3 and 4.2.4 reveals that ~30% of the solids in the latex and in the WRP were removed by buffer wash. These are most likely non-rubber constituents.

Table 4.2.3a – IAC40 latex solid content determination by freeze drying. Sample Liquid latex Wt. (g) Dried Wt. (g) Solids Content 091409_IAC40_L1 0.5009 0.1289 0.257 091409_IAC40_L2 0.5191 0.1289 0.248 091409_IAC40_L3 0.4981 0.1454 0.292 Average 0.266 Std Dev 0.023

Table 4.2.3b – IAC40 WRP solid content determination by freeze drying. Liquid WRP Wt. Sample (g) Dried Wt. (g) Solids Content 091609_IAC40_W1 0.5091 0.1222 0.240 091609_IAC40_W2 0.5111 0.1201 0.235 091609_IAC40_W3 0.5008 0.1217 0.243 Average 0.239 Std Dev 0.004

Table 4.2.4a – Solids content of IAC40 latex after washing the coagulated rubber. Sample Liquid latex Wt. (g) Washed/Dried Wt. (g) Solids Content 102709_IAC40_L1 0.5101 0.0986 0.193 102709_IAC40_L2 0.5019 0.0937 0.187 102709_IAC40_L3 0.4971 0.0955 0.192 Average 0.191 Std Dev 0.004

Table 4.2.4b – Solids content of IAC40 WRP after washing the coagulated rubber. Liquid WRP Wt. Sample (g) Washed/Dried Wt. (g) Solids Content 120909_IAC40_W1 0.5018 0.0898 0.179 120909_IAC40_W2 0.4997 0.0871 0.174 120909_IAC40_W3 0.5098 0.0916 0.180 Average 0.178 Std Dev 0.003

87 Table 4.2.5a and b show the solids content determined by precipitating the NR

latex and WRP into chilled methanol. Six vials of IAC 40 latex/WRP assays (three each),

were thawed to room temperature and directly precipitated into chilled methanol (4 oC).

The precipitated polymer was white. The polymer was then washed with buffer (100mM tris-HCl, pH = 7.5) and DI water to remove water-soluble solids and freeze-dried in the vacuum oven to constant weight.

Table 4.2.5a – Solids content of IAC40 latex obtained by precipitation in methanol. Liquid Latex Precipitated/Washed Sample Solids Content Wt. (g) /Dried Wt. (g) 071910_IAC40_L4 0.4468 0.0852 0.191 071910_IAC40_L5 0.3823 0.0721 0.189 071910_IAC40_L6 0.5482 0.1085 0.198 Average 0.192 Std Dev 0.005

Table 4.2.5b – Solids content of IAC40 WRP obtained by precipitation in methanol. Liquid WRP Wt. Precipitated/Washed Sample Solids Content (g) /Dried Wt. (g) 071710_IAC40_W4 0.6139 0.1023 0.167 071710_IAC40_W5 0.6993 0.1144 0.164 071710_IAC40_W6 0.7760 0.1217 0.157 Average 0.162 Std Dev 0.005

The solids content determined by methanol precipitation followed by buffer wash

are very similar, with the WRP solids being somewhat lower than the latex solids. Based

88 on these data, we considered the rubber content in the latex and in WRP 19 and 17 wt%, respectively. This may be the same within experimental error.

Utilizing the solids contents above, Table 4.2.6 summarizes the in vitro experiments carried out. The initial rubber weight was a calculated value evaluated from the liquid weight of the latex/WRP used multiplied by the solids content. The final dried/washed rubber weight is the final weight of the sample after 24 hours of incubation followed by washing with buffer and freeze-drying till constant weight. The mass difference between the final dried weight and the initial rubber weight is the mass gain, which is also shown as percentage value in Table 4.2.6. The final weight contained both gel and soluble fractions of NR. The sample identification code highlights the date, type of latex/WRP and the variation in the experiment. For instance,

KC_102009_IAC40_W_IP2 (10% EtOH) signified that the experiment was conducted on

10/20/2009 using WRP of IAC40 Hevea. The monomer used was IP and two samples had either 5% or 10% w/w ethanol added into the in vitro biosynthesis. With the exception of these two experiments, the other experiments were also monitored by in situ

Raman spectroscopy that will be discussed in Section 4.3; 80X and 50X refers to the size of the objective used in the Raman experiments. From Table 4.2.6, consistent mass gain was observed with the lowest being KC_111809_W_IP11 at 25%. The low mass gain of

25% from KC_111809_W_IP11 might be a result from loss of enzymatic activity because that particular batch of WRP was not assayed before use. Enzymatic activities are typically lost from repeated thawing and freezing cycles. The latex particles could coagulate when the water in the latex freezes. In all other cases, both the latex and the

WRP yielded 50% to 100% of mass gain.

89 Table 4.2.6 – Summary gravimetric data of in vitro NR samples. Initial Final Mass gain Mass gain Sample Rubber Dried/Washed (g) (%) Wt. (g) Rubber Wt. (g) KC_092909_IAC40_ 0.101 0.177 0.076 75 L_IP3 (80X) KC_092909_IAC40_ 0.098 0.179 0.081 83 L_IP4 (80X) KC_100909_IAC40_ 0.089 0.212 0.123 100(139) L_IP5 (80X) KC_102109_IAC40_ 0.090 0.193 0.103 100 (114) W_IP8 (50X) KC_110509_IAC40_ 0.090 0.134 0.045 50 W_IP9 (50X) KC_111209_IAC40_ 0.090 0.169 0.079 87 W_IP10 (50X) KC_111809_IAC40_ 0.104 0.130 0.026 25 W_IP11 (50X) KC_111909_IAC40_ 0.090 0.152 0.062 69 W_IP12 (50X) KC_101909_IAC40_ 0.090 0.137 0.047 52 W_IP1 (5% EtOH) KC_102009_IAC40_ 0.093 0.135 0.042 45 W_IP2 (10% EtOH)

The gel content was determined by freeze-drying insoluble and soluble portions to

constant weight after dialysis through 0.6 µm PTFE dialysis filter. Table 4.2.7

summarizes the results with the starting materials’ gel content in bold. The rubber

obtained from latex had higher gel content than that obtained from WRPs. This difference

arises from the gel being removed by the repeated centrifuging purification process in case of WRP extraction. From Table 4.2.5, it can be observed that gel content

consistently increased after incubation except for the samples where ethanol was added,

where the gel content remained the same within experimental error. This was expected

90 because ethanol was reported in the literature to break down the hydrogen bonding

present in the “soft” gel of NR.37

Table 4.2.7 – Gel content of in vitro NR samples.

Sample Gel content (%) KC_102709_IAC40_Latex 31.9 KC_092909_IAC40_L_IP3 (80X) 34.7 KC_092909_IAC40_L_IP4 (80X) 33.6 KC_100909_IAC40_L_IP5 (80X) 35.4 KC_120909_IAC40_WRP 23.8 KC_102109_IAC40_W_IP8 (50X) 29.5 KC_110509_IAC40_W_IP9 (50X) 29.2 KC_111209_IAC40_W_IP10 (50X) 32.9 KC_111809_IAC40_W_IP11 (50X) 27.1 KC_111909_IAC40_W_IP12 (50X) 25.9 KC_101909_IAC40_W_IP1 (5% EtOH) 21.8

KC_102009_IAC40_W_IP2 (10% EtOH) 23.0

Figure 4.2.2 shows the comparison between the SEC traces of the soluble

fractions obtained from IAC40 latex and IAC40 WRP. Figure 4.2.2 presents the zoomed

RI traces of the high and low MW regions for closer comparison. WRP has a shoulder

(H2*) at ~105 g/mol in the high MW region next to the main peak (H1) at ~106 g/mol.

This shoulder was not observed in the latex. In the low MW region, the WRP does not have L2, and the L3 peak is more pronounced in the WRP compared to latex.

Figure 4.2.2 – Zoomed SEC RI chromatograms of the soluble fractions from IAC40 and WRP for comparison: a) high MW region, b) low MW region.

The SEC traces show a substantial amount of low MW (< 4000 g/mol) fractions in both samples (Table 4.2.8). We have observed this in all samples we have analyzed.

The relative amounts were determined by the comparison of the RI traces, which are proportional to concentration. The SEC analysis shows that the soluble fraction obtained from WRP had slightly lower high MW components compared to the latex (24% vs

27%). Also, it was observed that all of the latex samples demonstrated an increase in the amount of high MW fraction with incubation of IP. The most pronounced was

KC_100909_L_IP5, in which the high MW fraction grew to 34% from the 27% of the latex. In the soluble fraction of the WRP samples, the high MW fraction also increased, but not as pronounced as in the latex examples. It should be noted here that the gel content also increased after incubation with IP in each case.

92 Table 4.2.8 – High and low MW parts of the soluble fractions obtained from the IAC40 latex and WRP. High MW fraction Low MW Sample (%) fraction (%) KC_102709_IAC40_Latex 27% 73% KC_092909_IAC40_L_IP3 (80X) 31% 69% KC_092909_IAC40_L_IP4 (80X) 30% 70% KC_100909_IAC40_L_IP5 (80X) 34% 66% KC_120909_IAC40_WRP 24% 76% KC_102109_IAC40_W_IP8 (50X) 25% 75% KC_110509_IAC40_W_IP9 (50X) 26% 74% KC_111209_IAC40_W_IP10 (50X) 24% 76% KC_111809_IAC40_W_IP11 (50X) 27% 73% KC_111909_IAC40_W_IP12 (50X) 29% 71% KC_101909_IAC40_W_IP1 (5% EtOH) 27% 73% KC_102009_IAC40_W_IP2 (10% EtOH) 29% 71%

Tables 4.2.9a and 4.2.9b summarize the SEC data. The H1 and H2 peak molecular weights were computed from the light scattering traces by ASTRA V. The H1 peak did not grow to higher MW after incubation with IP. In the soluble fraction obtained from the latex, a new peak appeared at ~ 105 g/mol, which was labeled as H2. Figure 4.2.3 shows the SEC RI trace of KC_092309_L_IP1 compared to the starting latex

(KC_102709_IAC40_Latex). All the latex samples show this new peak. However, in the case of the WRP samples, H2 overlaps with H2* (a shoulder peak in the starting WRP) and does not show a marked difference.

Figure 4.2.3 - SEC RI trace of KC_092309_L_IP1 and starting latex.

In the low MW region (not shown), the L3 and L4 peak intensities became more

pronounced after incubation with IP while the intensity of L1 decreased slightly

compared to the starting latex. (Similar behavior was observed for WRP samples.) L1

had a MW of approximately 400 g/mol while L3 and L4 had MWs of ~1,200 g/mol and

~2,000 g/mol respectively, using PIP calibration. Interestingly, the addition of EtOH introduced a new low MW peak labeled as L1*, which had a MW of ~200 g/mol. It is important to note that L1 always have the highest intensity after incubation with IP. ESI showed that L1was mostly composed of phospholipids, which were not involved in the in vitro NR biosynthesis.

95 H1 and H2 were integrated separately using Galactic GRAMS v5, a software

package that is able to deconvolute overlapping peaks and integrate the areas under the

respective curves in order to obtain the relative amounts based on the RI traces. It is

important to note that the Schultz-Zimm distribution is typically observed for

macromolecules; however, the software package only contains Gaussian fit, which was

used for this analysis. A modified built-in macro was used to fit Gaussian curves under

H1 and H2 peaks of high MW region of each SEC trace. The built-in macro used the least

square method; the modified macro applied the least absolute residual method, which was less sensitive to outliers (in this case, the H1 peak that overlaps with H2 peak, which skewed the least square fit to be broader, light green curve). (Figure 4.2.4)

Figure 4.2.4 - Difference between a) least square fit (unmodified) and b) least absolute residual fit (modified).

Figure 4.2.4 shows the difference between the least square fitting method and the least absolute residual method for KC_092309_L_IP1 as the two peaks are easily identified. It could be seen that the modified code fits better to the actual curve. It was important to note that the difference may seem very little in this example but was 96 significant in some other examples when a very broad H2 peak was present. The relative amounts were calculated as integral H2 / integral H1. Table 4.2.10 shows the summary.

When compared to gravimetric analysis, this data showed less NR growth. This is because this calculation considered the area of H2 as the new rubber. Furthermore, gravimetric analysis also included gel fractions while SEC data only analyzed soluble fractions. In short, although SEC showed less comparative growth than gravimetric analysis, the data was consistent and confirmed the new rubber growth as demonstrated by gravimetric analysis.

The 1H NMR spectrum of KC_092309_IAC40_L_IP1 in the regions of 0-3.5 ppm and 3.75-7.5 ppm are presented in Figures 4.2.5 and 4.2.6, respectively.

KC_092309_L_IP1 is a latex incubated with IP that showed 64% mass gain by gravimetric analysis and 100% growth by SEC. From Figure 4.2.5, it can be observed that the microstructure of NR is cis-1,4 polyisoprene.141 A small peak at 1.63 ppm might correspond to the trans initiator units as described in literature.41 The small signal at 0.9 ppm might indicate some cyclization.136

Figure 4.2.5 - 1H NMR (500 MHz) spectrum of KC_092309_L_IP1: 0-3.5 ppm region. (Concentration: 10 mg/mL, 128 scans, d1 = 10 sec, Pulse angle = 90o, T = 25oC, Solvent: toluene-D8.)

Figure 4.2.6 - 1H NMR (500 MHz) spectrum of KC_092309_L_IP1: 3.75-7.5 ppm region. (Concentration: 10 mg/mL, 128 scans, d1 = 10 sec, Pulse angle = 45o, T = 25oC, Solvent: toluene-D8.)

The broad peak of 5.69 ppm shown in Figure 4.2.9 above indicates 1,4-PIP chain enchainment (cis or trans conformation). The 3,4-enchainment was not observed since no 98 peaks could be observed at 4.4 to 4.8 ppm. The CH2=CH- proton of the vinyl group in

1,2-enchainment expected at 6.1 ppm was also not observed. The peaks obtained within

3.99 to 4.18 ppm might have arisen from alcohol or esters groups or from other

“abnormal” functional groups, which was previously reported to be present in NR.142

The 13C NMR spectrum of KC_092309_IAC40_L_IP1 in the regions of 0-50 ppm and 100-150 ppm are presented in Figures 4.2.7 and 4.2.8, respectively. In Figure 4.2.7, the characteristic peaks of trans-1,4 PIP at 16.0 ppm and at 26.9 ppm are not observed.143

In addition, the absence of peaks at 43.1 ppm and 40.4 ppm eliminates the 3,4-144 and

1,2-PIP145 microstructures, respectively. From Figure 4.2.8, the two peaks observed at

125.0 and 135.2 ppm correspond to cis-1,4 PIP.143 The characteristic peaks from other microstructures except cis-1,4 PIP are not observed, which is an indication that our in vitro NR is exclusively cis-1,4 PIP.

Figure 4.2.7 - 13C NMR (125 MHz) spectrum of KC_092309_L_IP1: 0-50 ppm region. (Concentration: 20 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

Figure 4.2.8 - 13C NMR (125 MHz) spectrum of KC_092309_L_IP1: 100-150 ppm region. (Concentration: 20 mg/mL, 10,000 scans, d1 = 10s, Pulse Angle = 90o, T = 25°C, Solvent: chloroform-D).

Figures 4.2.9 and 4.2.10 show the 13C NMR spectrum of KC_100909_L_IP5 in the 0-50 and 100-155 ppm regions, respectively. The peaks are better defined in this

spectrum because the NMR solvent was switched to benzene-D6; a study from Chen et

al.146 demonstrated that the separation between cis and trans signals of 1,4 polyisoprenes are the greatest in benzene-D6. The mass gain in this sample was 100% and the SEC

showed a ~60% growth of new NR. A 60-100% mass gain suggests that the new rubber

weight constitutes ~35-50% of the total mass of the sample. If the microstructure of the

newly formed rubber were other than that of cis-1,4 PIP, 13C NMR would be sensitive

enough to distinguish the other microstructures that may form during the incubation with

IP. In addition, if the polymerization mechanism were through free-radical or anionic

pathways, we should observe trans-1,4-, 3,4- and 1,2-PIP microstructures.143-145

100

Figure 4.2.9 - 13C NMR (125 MHz) spectrum of KC_100909_L_IP5: 0-50 ppm region. (Concentration: 50 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6).

Figure 4.2.10 - 13C NMR (125 MHz) spectrum of KC_100909_L_IP5: 100-155 ppm region. (Concentration: 50 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6).

101 The 13C NMR spectra of KC_102109_IAC40_W_IP8 are shown in Figures 4.2.11 and 4.2.12. This WRP sample had a mass gain of 100% and SEC growth of 50%. From the two spectra, only peaks corresponding to cis-1,4 PIP enchainment were observed, similar to in vitro samples that used raw IAC40 latex. In fact, all the samples had identical NMR spectra that only showed cis-1,4 PIP enchainment (attached in the

Appendix).

Figure 4.2.11 - 13C NMR (125 MHz) spectrum of KC_102109_W_IP8: 0-50 ppm (500 MHz Concentration: 60 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6).

102

Figure 4.2.12 - 13C NMR (125 MHz) spectrum of KC_102109_W_IP8: 95-155 ppm region. (Concentration: 60 mg/mL, 10,000 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6).

4.2.2. Incubation of Synthetic D-IP with Hevea Latex

In order to obtain spectroscopic evidence of the new rubber formation, deuterated

IP (D-IP) was incubated with IAC40 latex. The D-IP monomer (Source: Oak Ridge

National Laboratory (ORNL), TN, Received: 9/13/2010, Synthesized: 6/3/2008) was

sealed in a vacuumed ampule and claimed to be stable by ORNL. The purity of the

sample was determined prior to use. Figures 4.2.13 and 4.2.14 show the 1H NMR and

13C NMR spectra of IP (99%, Source: Sigma Aldrich) and D-IP obtained from ORNL, respectively. The synthetic procedure for the D-IP147 produces per-deuterated (i.e. 100%

deuterated) monomer; therefore, only little peaks corresponding to IP should appear in

the 1H NMR. The D-IP and IP NMR samples were prepared at the same concentration 103 and compared in Figure 4.2.13. The 1H NMR spectrum of D-IP (Figure 4.2.13) showed many peaks from unknown impurities and it cannot be confirmed that the compound was pure D-IP. However, it was worth noting that the D-IP and IP had the same characteristic odor.

Figure 4.2.13 - Comparison of 1H NMR spectra of IP (red) and D-IP (black). (300 MHz, Concentration: 20 mg/mL, 32 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

104 Since 1H NMR spectra cannot distinguish the structure of D-IP obtained from

ORNL, 13C NMR was taken and the comparison spectra are shown in Figure 4.2.14. In the 13C NMR spectrum, several peaks of D-IP resemble that of IP. When a compound is

Figure 4.2.14 - Comparison of the 13C NMR spectra of IP (red) and D-IP (black). (75 MHz, concentration: 20 mg/mL, 32 scans, d1 = 10s, Pulse angle = 90o, T = 25°C, Solvent: benzene-D6).

deuterated, the 13C peak is expected to split into multiple peaks and follow the

multiplicity rule of 2nI + 1, where n is the number of deuteriated nuclei attached to the

reference atom and I is the spin state of the attached deuteriated nuclei. The nuclear spin

of protons (H) is ½ whereas the nuclear spin of deuterium (D) is 1 which means that a 105 single deuterium nucleus can adopt three different spin states: 0, -1 and 1. For example, the 13C-D coupling in the 13C NMR spectrum of chloroform-D shows 2 x 1 x 1 + 1 = 3 multiplets. Since the 13C spectrum of the D-IP in Figure 4.2.17 shows multiple deuterium splitting, this confirmed that the sample obtained from ORNL contains per-deuterated IP with contaminants.

Gas chromatography (GC) data of IP from Sigma-Aldrich and Oak Ridge are shown in Figure 4.2.15. The major peak at 2.5 minutes elution time is assigned to the IP.

The small peak at 4.5 minutes for IP from is assigned to the p-tert-butylcatechol stabilizer. The ratio of the two peaks shows 99.1% purity for Sigmal-Aldrich IP. The D-

IP monomer shows a major peak at 2.5 min elution time as well as three other compounds (elution times: 0.9, 1.8, 3.2 min). Integration of the peaks from GC analysis of D-IP indicates 85.1% purity.

Figure 4.2.15 - GC chromatograms of IP (red) and D-IP (black).

106 The gravimetric results after incubation with D-IP are shown in Table 4.2.10.

(KC092910_L_IP2(50D-IP/50IP) stands for a mixture of 50% D-IP and 50% IP). The initial rubber weight was calculated based on 19.2 wt% solids content in the latex, determined by precipitating the latex into methanol and washing it with buffer. The NR

was freeze-dried till constant weight.

Table 4.2.10 – Gravimetric summary of D-IP experiments. Sample Initial Final Mass diff. Wt. of Wt. of Mass Rubber Dried/ b/w Final D-IP IP gain Wt. (g)* Washed Dried Wt. monomer monomer (%) Rubber and Ini. (g) (g) Wt. (g) Rubber Wt. (g) KC091610_L_ 0.086 0.098 0.012 0.09 0 14.0 IP1(100D-IP) KC091710_L_ 0.091 0.098 0.006 0.10 0 7.2 IP3(100D-IP) KC092910_L_ 0.098 0.122 0.024 0.05 0.05 24.3 IP1(50D-IP/50IP) KC092910_L_ 0.096 0.136 0.040 0.05 0.05 41.6 IP2(50D-IP/50IP) KC092910_L_ 0.097 0.160 0.062 0 0.10 64.1 IP3 (100IP control) KC092910_L_ 0.096 0.150 0.053 0 0.10 55.6 IP4 (100IP control) *based on 19.2 wt% rubber content

The experiments with 100% D-IP yielded 14.0% and 7.2% mass gain. In the experiments where 50/50 mixture of D-IP and IP were employed, more mass gains were observed (i.e. 24.3% and 41.6%). The control experiments (i.e. 100% IP) showed 64.1

and 55.6% mass gain.

107 Figure 4.2.16 shows the 13C NMR spectra of the experiments with 100% D-IP,

50/50 mixture of D-IP/IP and 100% IP in the 0-140 ppm and 20-35 ppm regions,

respectively. We observed the characteristic peaks of exclusively cis-1,4 PIP

microstructure in all samples. There was no peak splitting in any of the samples that

contained D-IP (the D-IP monomer displayed ~15 Hz, or 3~4 ppm peak splitting).

However, in the experiment with 100% D-IP, a new peak appeared at 30.55 ppm.

Currently, we are not able to identify this signal. It was also possible that the

contamination in the D-IP (i.e. 85.1% purity) terminated the active rubber chain ends in

the latex.

Figure 4.2.16 - 13C NMR spectra (125 MHz) of the D-IP experimental series, 091610_D- IP_latex 2 (top), 092910_L_IP1_50D-IP_50IP (middle) and 092910_L_100IP (bottom), o 0-140 ppm region. 10,000 scans, d1 = 10s, Pulse angle = 90 , Solvent: benzene-D6, Sample (concentration: 15 mg/mL for KC091610_L_IP1(100D-IP) and 50 mg/mL for both KC092910 experiments.)

Latex and WRP were also incubated with 13C labeled IP. The NMR spectra showed no

13C signals in the rubber.

In summary, the addition of synthetic IP to in vitro NR biosynthesis increased the

solids content in WRPs and latex, without incorporating into the rubber.

108 4.2.3. In Vitro NR Biosynthesis under CO2 Atmosphere

In plants, the mitochondrial metabolism is controlled by the synthesis of adenosine triphosphate (ATP). A process called tricarboxylic acid (TCA) cycle, which releases carbon dioxide is a fundamental component of the ATP synthesis.82 Therefore, the atmosphere within the plant medium, i.e. cytosol, is enriched by dissolved CO2. In order to closely mimic the environment where NR biosynthesis occurs in vivo, we have developed a series of experiments where the closed system consists of only CO2 atmosphere. (Chapter 3.2.2.4: Figure 3.2.3)

Under CO2 atmosphere, the reaction mixture of WRP and isoprene monomer had a slightly yellower tint compared to experiments conducted under ambient condition.

Please note that “ambient condition” refers to normal atmospheric conditions, where the vials were not purged with carbon dioxide or any other gas.

The gravimetric analysis and gel content determination of four CO2 experiments are presented in Table 4.2.11 and Table 4.2.12, respectively (the Raman monitoring of these experiments will be discussed in Section 4.3). Note that the initial rubber weight was taken based on the 19.2 % solid (i.e. rubber) content in the latex. All the WRP experiments showed significant mass gain, with KC121409_W_IP3 (CO2/50X) showing the highest mass gain at 98%. The gel content of the reaction performed utilizing IAC40

WRP under carbon dioxide atmosphere was measured only for one sample, which did not seem to change after incubation. This is different than what we have previously observed under normal atmospheric conditions, where the gel content was found to increase after

109 incubation. For the experiment conducted with IAC40 latex, it was found that there was minimal mass gain (i.e. 7%) and the gel content remained unchanged.

Table 4.2.11 - Gravimetric analysis of in vitro NR biosynthesis under CO2 atmosphere. Mass diff. Final b/w Final IP Initial Mass Latex/WRP Dried/Washed Dried Wt. Sample monomer Rubber gain Wt. (g) Rubber Wt. and Ini. Wt. (g) Wt. (g) (%) (g) Rubber Wt. (g) KC_120809_L_I 0.6406* 0.130 0.123 0.131 0.008 7 IP1 (CO2/50X) KC_121109_W_ 0.4697 0.100 0.090 0.136 0.046 51 IP1 (CO2/50X) KC_121109_W_ 0.6715* 0.133 0.129 0.208 0.079 61 IP2 (CO2/50X) KC_121409_W_ 0.5085 0.102 0.096 0.190 0.094 98 IP3 (CO2/50X) *Latex/WRP weight for KC_120809_L_IP1(CO2/50X) and KC_121109_W_IP2 (CO2/50X) are higher than the other two samples due to Raman measurement configurational needs, as described in the experimental section.

Table 4.2.12 - Gel content of in vitro NR samples. Sample Gel content (%) KC_102709_IAC40_Latex 31.9

KC_120809_IAC40_L_IP1(CO2/50X) 32.4 KC_120909_IAC40_WRP 23.8

KC_121109_IAC40_W_IP1(CO2/50X) -*

KC_121109_IAC40_W_IP2(CO2/50X) -*

KC_121409_IAC40_W_IP13(CO2/50X) 23.3 *not available

However, despite minimal mass gain after the incubation, the SEC trace, shown in

Figure 4.2.17 shows the appearance of a peak at ~105 g/mol, which we have assigned to the formation of new rubber. Figure 4.2.18 shows an example of IP incubation with

110 WRP. In the low MW region, while no change was observed for the experiment

conducted with latex (i.e. KC120809_L_IP1(CO2/50X), a new peak L2 was observed for

the experiment conducted with WRP (KC121409_L_IP3(CO2/50X). The SEC data are

summarized in Table 4.2.13.

Figure 4.2.17 - SEC comparison RI traces of KC_102709_Latex_IAC40 and KC_121109_L_IP1(CO2/50X).

Figure 4.2.18 - SEC comparison RI traces of KC120909_IAC40_WRP and KC121409_W_IP3 (CO2/50X).

111 The other traces obtained with WRP (KC_121109_IAC40_W_IP1(CO2/50X and

KC_121109_IAC40_W_IP2(CO2/50X) were similar (not shown).

Table 4.2.13 - SEC analysis of the soluble fractions obtained from IAC40 latex/WRP

before and after incubation with IP in CO2. H1 peak H2 peak H1 H2 L4 L3 L2 L1 Sample ID MW MW (min*) (min*) (min*) (min*) (min*) (min*) (g/mol) (g/mol) 102709_IAC 36.15 1.6x106 - - 55.90 58.08 59.47 61.50 40_latex 120809_IAC 40_L_IP1 36.08 1.6x106 41.35 1.5x105 55.16 57.91 59.02 60.88 (CO2/50X) 120909_IAC 40_WRP 35.93 2.0x106 - - 55.11 58.16 - 61.62 (CO2/50X) 121409_IAC 40_W_IP3 36.21 1.5x106 40.84 4.3x105 55.08 58.53 59.81 62.03 (CO2/50X) *elution time

4.2.4. In Vitro NR Biosynthesis in the Presence of Amylene

Control experiments with the addition of amylene (Figure 4.2.19), an isomer of

IP, were carried out. Of the four reactions, three were repeats of previous experimental

conditions.

Figure 4.2.19 – Chemical structure of amylene.

112 One reaction was monitored by Raman spectroscopy which will be discussed in

Section 4.3. The gravimetric results are summarized in Table 4.2.14. Limited mass gains

were observed in these samples. The SEC for KC_101510_L_Amy1 was prepared by

dissolving the product in benzene and the gel fraction was filtered out using a 0.45 μm

PTFE membrane filter. From Figure 4.2.20 below, the SEC of

KC_101510_L_Amy1showed no changes in the RI trace. No new peaks were

Table 4.2.14 – Gravimetric summary for Amylene experiments. Final Mass diff. b/w Initial Mass Dried/Washed Final Dried Wt. Sample Rubber Wt. gain Rubber Wt. and Ini. Rubber (g) (%) (g) Wt. (g) KC101510_L_Amy1 0.0976 0.1118 0.0142 14.6% KC101510_L_Amy2 0.0961 0.1211 0.0250 26.1% KC101510_L_Amy3 0.0967 0.1114 0.0147 15.1%

Figure 4.2.20 – SEC comparison RI traces of 102709_Latex and KC_101510_L_Amy1 observed after incubation of IAC40 NR latex with amylene. The other two experiments (KC101510_L_Amy2 and KC101510_L_Amy3, not shown) showed similar results with KC101510_L_Amy1 with no changes in the SEC traces. 113 Based on the experiments presented in Section 4.2, we can conclude that the addition of synthetic IP to NR latex and WRP results in net mass gain, but the IP does not incorporate into the rubber. It is very likely that the IP triggers a biological mechanism that releases IPP, leading to new rubber formation.

4.3. In Situ Micro-Raman Monitoring of In Vitro Natural Rubber (NR) Biosynthesis

In order to monitor IP incorporation into NR in situ, a new Raman methodology had to be developed. This was accomplished in collaboration with Professor Sokolov and his Ph. D. student Andrei Malkovskiy in the Department of Polymer Science at the

University of Akron. The method developed is detailed below.

4.3.1. Raman Monitoring using Glass Slides

Spectra were separately obtained for the synthetic high cis-PIP (prepared by anionic polymerization), the IP stabilized by 100 ppm p-tert-butylcatechol (Sigma-

Aldrich) and a WRP (RRIM600, USDA) sample (KC_030209_WRP1). 10 mg of the

WRP sample was placed between two fresh pre-cleaned microscope glass slides

(Fisherbrand® plain microscope slides, 25x75x1 mm) and positioned perpendicular to the incident laser beam (λ = 647 nm) of moderate intensity (~ 1mW). RRIM600 WRP samples taken from a refrigerator a few minutes prior to the measurements, in which the

WRP should be still active, were used in all the measurements. The slides were pressed tightly in a standard spring holder. The spectrum of the sample was measured together

114 with the corresponding background spectra of both the top and the bottom glass slides.

All the data were corrected by subtracting the background glass signals from the spectra of the samples of interest. We found many characteristic peaks of PIP: two peaks at

~1000 and ~1040 cm-1, three broad peaks at ~1320, ~1370 and ~1450 cm-1 and a peak at

~1670 cm-1. However, after a detailed analysis on the combination of spectra, we decided

to focus only on the peak at 1670 cm-1, because of its sharpness along with the highest intensity of absorption. This peak is attributed to the C=C bond of PIP. We have found

this signal in both the synthetic PIP and the RRIM600 WRP. The shoulder next to this

peak in the PIP standard remains unidentified at this point. The signal of the conjugated double bond in IP appeared at 1640 cm-1 (see Figure 4.3.1).

Figure 4.3.1 - Signals attributed to the C=C Raman-active vibrations in PIP and IP (Isoprene: liquid IP with stabilizer; Mixed sample: WRP RRIM600+IP 2/1 (w/w)).

115 The difference in the vibrational energy for the IP and PIP peaks is less than 30

cm-1, which was easy to resolve with Sokolov’s Raman system. Figure 4.3.1 also shows

the signal of the sample marked “mixed” in which IP was added to the RRIM600 WRP

(2:1 w/w WRP:IP) and allowed to react for ~5 minutes. In this case, the spectrum of the

bottom glass slide was not measured because the sample was opaque. Six different

spectra were measured for the mixed sample: 2 spectra from random spots, 3 spectra

from different WRP particles and 1 spectrum from the last latex particle exposed to five

minutes of laser illumination (to observe whether there was sample damage from the

laser). In the mixed sample, the signals of both IP and PIP were detected, with the PIP signal being less intense.

4.3.2. Raman Monitoring with Micro-cavity Slides.

IP has a boiling point 34 oC and easily evaporates at room temperature. To avoid

IP evaporation between two glass slides, special glass slides (Pearl®, clear glass with ground edges, 25.4x76.2 mm, ~1.1 mm thick) with a small cavity have been purchased.

The top slide was Pearl® cover glass slides (~0.15 mm thickness). The bottom slide had a cavity, which allowed higher volumes of material to be placed on the slide, while at the same time ensuring a better seal of the two-slide assembly to avoid IP evaporation. The latex invariably adhered to the outer edge of the cavity when sealed due to strong capillary forces. This made signal acquisition possible even when the cavity was only partially filled and limited the evaporation of IP. The Raman system also allowed video- monitoring of the sample under the microscope. It enabled observation of the probed

116 volume and the exact spot illuminated by the laser. An external white lamp was

connected to the microscope to provide a more homogeneous illumination source. With

the help of this camera, another observation was made: the spectra measured at the

position of a single latex particle were generally much stronger than those measured

away from any particles, while the fluorescence background from the bottom glass slide

was also weaker. This probably indicates that the observed Raman signal comes mostly

from the particles and not from the surrounding media. Despite much better sealing of the

sample between the glass slides with a cavity, IP evaporation could not be excluded.

4.3.3. Raman Monitoring in Sealed Silanized Vials

It is crucial to develop Raman experiments methodologies where we can be sure that no IP is lost due to evaporation. 0.5 mL IAC40 WRP and IAC 40 latex samples were sealed in glass vials (Fischer Scientific, 1ml silanized glass vials) using a crimper. After the injection of IP (0.102g, 0.15mL, 1.50mmol) through the hermetic cap of the crimp, the contents of every vial were vigorously shaken for a minute to ensure good mixing of the IP and the initial rubber particles. Then the bottle was fixed under a long-working distance Mitutoyo APO SL50 objective (50X, NA = 0.42) in horizontal position.

Photographs of the setup are shown in Chapter 3.2.2.3 (Figure 3.2.2). Measurements were also performed with 20X and 80X (NA = 0.42) Mitutoyo long-working distance objectives as well. However, the 20X objective was collecting significant extra Raman signals from the glass walls of the vial (due to larger focal volume of the objective). The

80X objective gave much lower overall signal intensity, when compared to the 50X 117 objective. After comparing the data measured with the three objectives described above, the 50X objective was chosen as the one providing the best Raman signal of IP and PIP with very low glass background signal. The Raman signals were measured immediately after the IP injection and mixing (about 1 minute) through the glass walls. The spectra were collected for five minutes for every data point with intervals of five minutes between the measurements for the duration of the first six hours of the experiment. Then the samples were incubated for 18 more hours for a total incubation time of 24 hours.

Figure 4.3.2 shows the positioning of the laser beam. It probes an area of ~1 cubic micron, which is roughly the size of an average particle and very localized. The

RRIM600 average particle size is approximately 1.1±0.6 μm and the IAC40 average particle size is approximately 1.6±0.5 μm. This makes the quantification of the Raman data difficult.

Figure 4.3.2 - The schematics of the bottle illuminated by the Raman beam.

Another challenge was the strong fluorescence. Continuous illumination of a single spot raised the fluorescence of the sample to a degree that the experiment could not be continued. A different spot had to be picked and re-focused, which makes quantitative 118 analysis impossible. This problem is illustrated in the Figure 4.3.3 for an IAC40 latex

sample; however the fluorescence problem also persisted for the WRP samples. We speculated that one reason contributing to the scattering of data points might be the 80X objective, which focuses on a small area of the rubber particles. Therefore, we decided to enlarge the probing area by switching to a 50X objective and also by blocking the beam between collections, which was applied in all subsequent experiments.

Figure 4.3.3 - Fluorescence problems illustrated for KC_092309_L_IP1 with 80X objective. (092309_L_WRP: 0.5340g IAC40 latex, 0.1022g IP, 24h, RT).

Figure 4.3.4 presents a set of Raman spectra of an experiment with IAC40 WRP

(KC_102109_W_IP8 (50X)) as a function of time, where the IP and PIP Raman signals are marked. It is apparent that the IP peak intensity decreases while the PIP signal intensity increases. The experiment was repeated twice. For more quantitative analysis, the spectra in the frequency range of interest were fitted by two Lorentzian peaks, one for the IP signal at 1640 cm-1 and another for PIP signal at 1670 cm-1 (Figure 4.3.4).

119

Figure 4.3.4 - Raw spectra for a sequence of data points of KC_102109_W_IP8 (KC_102309_WRP_IP8: 10 min between each spectra, 0.5079g IAC40 WRP, 0.1022g IP, 24h, RT).

Due to noise in the spectra, the fit gives fluctuating values of the peak positions

and widths. In order to improve the accuracy of our analysis, we calculated the average

values of these parameters for each data set. In the next iteration, the spectra were fitted

again with the peak width and position fixed at the averaged values of the set. The areas

under the fit curves were used to estimate the integrated intensity of each signal. The ratio of the PIP peak area to the sum of the IP and PIP peak areas thus is a semi- quantitative illustration of the kinetics of PIP formation. Absolute measurements of the

Raman intensity revealed sometimes strong change of the signal due to the fact that the overall amount of material next to the top wall of the vial (that is placed on its side under the microscope objective) might have changed over time due to gravitational forces, preventing us from plotting just the growth in the PIP peak area.

Figure 4.3.5 presents a comparison of the (/ ) – time plots for two

representative experiments. In both cases, we see a decrease in the early part of the 120 experiment. KC_120209_WRP_IP13 remained constant while KC_110509_WRP_IP9

showed an increase. After extensive experimentation, we realized that this was due to an artifact as a consequence of inadequate mixing. We also observed that PIP formation was seen only when a yellow color appeared in the mixtures. This was the case for

110509_WRP_IP9 (red circles) showing an increasing trend, most likely due to adequate mixing. We believe that the yellow color indicates the presence of ionic species.

120209_WRP_IP13 (green stars) was white (no yellow color appeared upon addition of the IP). The data points are from single measurements over time.

Figure 4.3.5 - Normalized PIP formation plots for two repeat experiments using 50X objective. (KC_110509_WRP_IP9: 0.5028g WRP, 0.1022g IP, 24h, RT. KC_120209_WRP_IP13: 0.5097g WRP, 0.1021g IP, 24h, RT).

Figure 4.3.6 shows PIP formation plots for two samples that changed their color to yellow upon IP addition, observed under the 50X objective, and the beam was blocked in between data acquisition steps. The absence of the initial decrease, as in Figure 4.3.6, 121 was the evidence for good sample mixing. The intensity of the PIP signal doubled in the two samples (102109_W_IP8 and 111209_W_IP10). The size of the error bars showed the reliability of the Lorentzian fits and consistency between the experimental points.

Figure 4.3.6 - Normalized PIP formation plots for two repeat experiments with 50X objective with error bars. (KC_102309_WRP_IP8: 0.5079g WRP, 0.1022g IP, 24h, RT. KC_111209_WRP_IP10: 0.5843g WRP, 0.1022g IP, 24h, RT).

Figure 4.3.7 shows IP/(PIP+IP)–time plots. Qualitatively, the plots demonstrated decreasing trends. The IP signal never disappeared completely. This could be explained by the position of the probed volume. The Raman spot was probing a very small volume, located next to the glass wall of the silanized vial, where IP could get trapped in the confined space.

122

Figure 4.3.7 - Normalized IP consumption plots for three repeat experiments with 50X objective. (103209_WRP_IP8: 0.5079g WRP, 0.1022g IP, 24h, RT. 110509_WRP_IP9: 0.5028g WRP, 0.1022g IP, 24h, RT. 111209_WRP_IP10: 0.5843g WRP, 0.1022g IP, 24h, RT).

Figure 4.3.8 shows the result of in situ Raman monitoring experimental method

that had been optimized after many experiments. The experimental procedure was as

follows: 0.5 g active IAC40 WRP or IAC40 latex were sealed in glass vials (Fischer

Scientific, 1ml silanized glass vials) using a crimper. After the injection of isoprene

(0.102 g, 0.15 mL, 1.50 mmol) through the hermetic cap, the contents were vigorously shaken for one minute to ensure complete mixing. The color of the mixture was observed and noted. The reactions were monitored by in situ Raman spectroscopy to observe PIP growth. The sealed polymerization vial was fixed under a long-working distance 50X objective from Mitutoyo with NA = 0.42. The laser source was Lexel RamanIon Krypton laser and the excitation wavelength was set at 647 nm. Raman signals were collected by a

123 Horiba Jobin-Yvon Labram HR single monochromator equipped with a nitrogen-cooled

CCD camera through the glass walls of the vials. Spectra were collected for five minutes for every data point, and then the laser was blocked for five minutes to avoid heating and fluorescence. This sequence was repeated for the duration of the first six hours of the experiment. The samples were incubated for a total of 24 hours. From Figure 4.3.8, the

Raman plots demonstrate IP decrease and PIP increase using two equations: /

and / . We are currently in the process of trying to quantify the data

based on gravimetric measurements, however in relative terms we found that the change

in normalized Raman intensity from the beginning to the end of the experiment is +0.1

unit for PIP and -0.1 unit for IP.

Figure 4.3.8 - Raman monitoring of• in vitro NR biosynthesis. (KC_102109_WRP_IP8: 0.5079 g IAC40 WRP, 0.1021 g IP, 24 hrs, RT).

In summary, in situ Raman monitoring showed IP depletion and PIP growth.

4.3.4. Raman Monitoring of Incubation with Deuterated-Isoprene (D-IP)

124 In Chapter 4.2, we attempted to observe the incorporation of IP into NR by means of D-IP. The D-IP incubation with NR was also monitored by Raman spectroscopy.

Figure 4.3.9 shows the Raman monitoring of KC091710_L_IP3 (100DIP). The D-IP has a characteristic peak at 1590 cm-1 whereas IP, as determined from previous experiments, has a peak at 1630 cm-1. The in situ Raman monitoring showed some scatter but no overall change in the / – time plot was observed.

Figure 4.3.9 - Raman monitoring of KC091710_L_IP3 (100DIP).

4.3.5. Raman Monitoring under CO2 Atmosphere

Figure 4.3.10 shows the in situ Raman monitoring of KC_121409_W_IP3

(CO2/50X), representing the experiment utilizing WRP and conducted under carbon dioxide atmosphere (see Section 4.2 for the gravimetric analysis). When compared with previous WRP experiments

125

Figure 4.3.10 - PIP growth comparison between WRP experiments and KC_121409_W_IP3 (CO2/50X) 0.5085 g of IAC40 WRP, 0.102 g of IP, 24 hrs under o CO2 atmosphere, at 25 C.

The reaction mixture with IAC40 latex under CO2 atmosphere had a stronger yellow tint in comparison with the sample subjected to atmospheric conditions. Figure

4.3.11 compares the in situ Raman monitoring of KC_120809_L_IP1(CO2/50X),

experiment (conducted in CO2 atmosphere) and KC_092909_L_IP4, the experiment

conducted under ambient conditions. The rate of polymerization and/or enzymatic

activity does not appear to be affected by CO2.

126

Figure 4.3.11 - PIP growth comparison between KC_092909_L_IP4 and KC_120809_L_IP1(CO2/50X), 0.6406 g of IAC40 Latex, 0.130 g of IP, 24 hrs under o CO2 atmosphere at 25 C.

In general, from the Raman spectra in Figures 4.3.10 and 4.3.11, the rate of initial

NR growth is higher for experiments conducted with latex than WRPs, as observed from

the PIP/(IP+PIP)t≈0 min ratio of ~0.150 for atmospheric IAC40 latex and much lower (i.e.

~0.075) for IAC40 WRP (Figure 4.3.12).

4.3.6. Effect of IP Addition on Latex Particle Size

The effect of IP addition to NR latex particle size was investigated using

enzymatically inactive ammonia-stabilized latex (Neotex HA (High Ammonia Content))

of Hevea. Figure 4.3.12 shows an image of Hevea latex particles in focus. The

distribution of particle size of about 300 particles measured manually by ImageJ (image

processing program) appeared to be close to a Gaussian distribution (Figure 4.3.2).

127

Figure 4.3.12 - Optical images of Hevea latex particles (Neotex HA) casted on a flat glass slide.

F Gauss fit of Data1_F 100 Total # particles measured = 325

80 center ~1,1 micron

20 Frequency of appearance, units of appearance, Frequency

0 0.6 0.8 1.0 1.2 1.4 1.6 1.8 2.0 2.2 Particle diameter, mkm

Figure 4.3.13 - Size distribution of Neotex HA latex particles.

The average particle size of Neotex HA was found to be around 1.1 µm, which was similar to the RRIM600 Hevea clones obtained from the USDA (size = 1.1 µm) but smaller than IAC40 clones which had a size of 1.6 µm. The addition of IP (0.l mL, 0.1 mmol) to 0.2 mL (0.19 g) of Neotex HA latex led to significant shrinking of the latex particles to a size of approximately ~250 nm, four times smaller than their original size of

128 ~1 µm. Similar observation was reported from the USDA for raw Hevea latex.148 After

24 hours, the Neotex HA latex coagulated; however, the USDA reported that the particles grew back to their original size after 24 hours.

In summary, in situ Raman monitoring gave additional insight into the effect of adding synthetic IP to enzymatically active Hevea latex and WRP. However, the source of the mass gain and changes in the SEC traces remains unexplained. The Puskas group is currently collaborating with the CNRS (France) and (Spain) within the framework of an

IUPAC-NSF project to investigate terpenoid biosynthesis and shed light to the unexplained phenomena presented in this thesis.

4.4. Macroinitiator Synthesis

The synthesis of oligoisobutylene-neryl pyrophosphate macroinitiators will be described in this chapter. Synthesis of a PIB-based macroinitiator was previously proposed by Gautriaud and Puskas to yield a PIB-NR copolymer.149 Ms Gautriaud

completed and optimized the first two steps of the macroinitiator synthesis. For step 1,

she completed the synthesis with 90% conversion. In step 2 (allylic oxidation), she achieved 35% conversion. She proceeded to perform a number of model compound reactions. Based on her findings, the synthetic scheme was revised and will be presented in this thesis. A modified synthetic strategy was followed as shown in Figure 4.4.1.

129 PPTs DHP

CH2Cl2 OH 27°C, 4h OO 1 step 1 2 Nerol PN

SeO2 OH salicylic acid

CH2Cl2 27°C, 14h OO step 2 3 PN-OH

OTs TsCl

N,N-dimethylbutylamine K2CO3/H2O OO pH=10 4 step 3 PN-Ts

130 NaH OPIB PIB-OH

THF OO 90°C, 5d, N2 step 4 5 PIB-PN

PPTs OPIB IPA(2):Hx(1)

RT, 20h step 5 OH 6 PIB-Nerol

NBS, S(CH2)2 OPIB

-40oC, 1h CH2Cl2 Br step 6 7 PIB-Nerol-Br

OPIB DOWEX AG50X8 O O

(n-Bu4N)3HP2O7:MQ water O P O P OH step 7 8 OH OH PIB-NPP

Figure 4.4.1 - Synthetic strategy to produce PIB-NPP macroinitiator. PPTs = pyridinium p-toluenesulfonate; DHP = dihydropyran; NBS = N- bromosuccinimide; TsCl = tosyl chloride; IPA = isopropyl alcohol. with PIB-O =

CH3 CH3 CH3

C CH2 C CH2 C Cl n CH2 CH3 CH3 O 131

In step 1, the hydroxyl group of the commercially available starting compound,

nerol, was protected with tetrahydropyran (THP) to form a nerol tetrahydropyranyl ether

(product 2, PN). The PN was oxidized at the C8 position with selenium dioxide (SeO2)

(step 2) to yield the corresponding allylic alcohol (product 3, PN-OH). Subsequently, the hydroxyl group was functionalized with a tosyl group (step 3) to yield an allylic tosylate

(product 4, PN-OTs). In step 4, nucleophilic substitution of the tosyl group in PN-OTs with a PIB carrying a primary hydroxyl head group in the presence of sodium hydride

(NaH) to yield product 5, PIB-PN. The tetrahydropyranyl ether protecting group in PN-

PIB was then deprotected to produce the corresponding alcohol (step 5, product 6, PIB-

Nerol) which in turn was further functionalized with an allylic bromide using the Corey-

Kim reagent (step 6) to yield product 7, PIB-Nerol-Br. Finally, PIB-Nerol-Br was functionalized with a pyrophosphate end group using (Bu4N)3HP2O7 (product 8, PIB-

NPP) to yield our final product of PIB-NPP.

4.4.1. Synthesis of Protected Nerol (PN, Product 2)

The 1H NMR spectrum of the starting compound (nerol) is shown in Figure 4.4.2.

The two resonances at (f) (5.06 ppm, t, 1H, J = 27 Hz) and (b) (5.43 ppm, t, 1H, J = 27

Hz) ppm correspond to the methine protons with triplet splitting patterns. They are split by the two neighboring protons of the CH2 (a, e) to yield a triplet pattern. The protons of

the CH2 group (a) (d, 2H, J = 12 Hz) next to the oxygen show up as a doublet at 4.07 ppm. The four protons of the two CH2 groups, (d) and (e), appear at 2.02-2.13 with

132 integral of 4.34 relative to (b). Finally the 3 CH3 groups (g’), (h), and (c’) are identified as singlets at 1.59, 1.67 and 1.72 ppm, respectively, with an integral of 9.20.

1 Figure 4.4.2 - H NMR spectrum of nerol. (solvent: CDCl3, nt = 32, d1 = 1s, AQ = 2s, 300 MHz, Appendix: B.4.4.2)

After the nerol react with 3,4-dihydropyan, the residue was purified by flash chromatography (eluent: ethyl acetate/hexane, 1:8 v/v; TLC: Rf = 0.7) on silica gel to yield PN (~90%). Figure 4.4.3 shows the 1H NMR spectrum of PN. The two resonances

(f) and (b) corresponding to the methyne protons are found in 5~5.5 ppm. The two protons at (a) (4.12 ppm, d, 2H) shifted downfield. New peaks corresponding to the

dihydropyran protecting group are found. The peak at 4.6 ppm and is assigned to the

proton (i) (4.61 ppm, t. 1H, J = 9 Hz) with an integral of 1.08. The doublet of multiplets

at 3.50 and 3.90 ppm corresponds to the two hydrogens of the CH2 group on (j) (m, 2H) next to the oxygen of the pyran group. The remaining six protons of the protecting pyran

133 ring, (k), (l) and (m), appear as multiplets in the aliphatic region and overlaps with the

CH3 groups [(c’), (g’), (h)] of nerol. The integral of the aliphatic region (1.4~1.8 ppm) is

15.42, which agrees closely with the 15 protons (6 from (k), (l), (m) and 9 from (c’), (g’),

(h)) present in PN.

1 Figure 4.4.3 - H NMR spectrum of PN (product 2). (solvent: CDCl3, nt = 64, d1 = 1s, 300MHz, AQ = 2s, Appendix: B.4.4.3).

The 13C NMR spectrum further confirmed the chemical structure derived from the

1H NMR spectrum. In Figure 4.4.4, the number of carbon peak resonances matched with

the compound. The four carbons with double bonds, (B), (C), (F), and (G), appear at

120~140 ppm. The resonance at 97 ppm corresponds to the carbon (I) between the oxygen atom of PN and the oxygen atom of the pyran ring. The resonances at 62 and 64 ppm correspond to the carbon atom next to the oxygen in the PN moiety (A) and (J). CH3

134 carbons resonances of (G’), and (C’) appear at δ = 18, and 26 ppm, respectively. The

CH2 carbon resonances of (D) and (E) appear at δ = 33 and 28 ppm. The carbon resonances of the pyran ring [δ = 31 ppm (M) and δ = 19 ppm (L), and δ = 30 ppm (K)] confirmed the protection of nerol.

13 Figure 4.4.4 - C NMR spectrum of PN (product 2). (solvent: CDCl3, nt = 5,000, d1 = 1s, 125 MHz, AQ = 1.3s, Appendix: B.4.4.4).

4.4.2. Synthesis of Protected Nerol-OH (PN-OH, Product 3)

PN was oxidized to yield PN-OH for the eventual attachment of PIB. Initially, the conversion of PN to PN-OH was very low (~10%). The literature and Ms. Gautriaud reported 35-40% conversion. The problem of low conversion was solved by washing the peroxide with chloroform to remove the water and titrating t-butyl hydroperoxide with potassium manganite (KMNO4) solution to determine the peroxide concentration. After

135 24 hours, diethyl ether was added and the mixture was washed with saturated sodium

hydrogen carbonate aqueous solution. The compound was extracted with anhydrous

diethyl ether, dried over MgSO4, filtered, and concentrated. The crude oil was purified by flash chromatography (eluent: ethyl acetate/hexane, 1:5 v/v; TLC: Rf = 0.15) on silica gel to yield PN-OH. This method yielded ~45% conversion.

1 Figure 4.4.5 shows the H NMR spectrum of PN-OH. The CH3 of (h) from nerol

(δ = 1.6 ppm) is oxidized to CH2-OH and shifted downfield to δ = 3.95 ppm with an integral of 2.17. The integral of 1.4~1.8 ppm is now 12.06 (6 from (k), (l), (m) and 6 from (g’), (c’)), which agrees right the transformation. The CH proton of (f) shifted downfield to 5.36 ppm, confirming the allylic oxidation of (h) (3.92 ppm, d, 1H).

1 Figure 4.4.5 - H NMR spectrum of PN-OH (product 3). (solvent: CDCl3, nt = 64, d1 = 1s, 300 MHz, AQ = 2s, Appendix: B.4.4.5).

136 The 13C NMR shown in Figure 4.4.6 verifies that the allylic oxidation was successful. (H) shifted from 24 to 68 ppm, corresponding to the CH2 carbon next to the -

OH group.

13 Figure 4.4.6 - C NMR spectrum of PN-OH (product 3). (solvent: CDCl3, nt = 5,000, d1 = 1s, 125 MHz, AQ = 1.3s, Appendix: B.4.4.6).

4.4.3. Synthesis of Protected Nerol Tosylate (PN-Ts, Product 4)

The alcohol leaving group of PN-OH was tosylated to yield PN-Ts, which is a more reactive leaving group. Ms Gautriaud previously attempted a method introduced by

Grubbs et al., but resulted in low conversion. Instead of the traditional 4- dimethylaminopyridine (DMAP) catalyst, a “greener” method to yield tosylates from allylic alcohols was developed by Morita et al. N,N-dimethylbutylamine was used as the catalyst in suspension with water in basic conditions (pH = 10). The excess tosyl chloride

137 was quenched with N,N-dimethylethylenediamine and become water-soluble for workup.

The crude product was purified by flash chromatography on silica gel to yield PN-OTs

(12.62 g, 92%): TLC (hexane/ether, 15:1, v/v, Rf 0.5.)

1 Figure 4.4.7 shows the H NMR spectrum of PN-Ts. CH2 protons (h) (4.1 ppm, s,

2H) shifted downfield compared to the (h) of allylic alcohol at 3.9 ppm. New resonances attributed to the tosyl group ((n) (δ = 7.87 ppm, d, H), (o) (δ = 7.25 ppm, d, H) and (p) (δ

= 2.42 ppm, s, H) appeared with the integration ratios of (n):(o):(p) as 1.98:2.00:2.99.

Figure 4.4.7 - 1H NMR spectrum of PN-Ts (product 4). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.7).

Figure 4.4.8 shows the 13C NMR spectrum of PN-Ts. Carbon resonance (D’) of

the hydroxyl group at δ = 69 ppm in the PN-OH shifted downfield to δ = 71 ppm in PN-

OTs. New carbon resonances corresponding to the tosyl group (δ = 128 (N), δ = 132 (O)

138 δ = 138 (N’), δ = 144 (O’) and δ = 36 (P)) appeared at the expected positions, confirming the structure of the product.

Figure 4.4.8 – 13C NMR spectrum of PN-Ts (product 4). (125 MHz, 5,000 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, AQ = 1.3s, Appendix: B.4.4.8).

4.4.4. Synthesis of PIB-PN (Product 5)

One of the greatest challenges faced in this project was the attchment of the model nerol compound to primary hydroxyl-PIB. The primary hydroxyl-PIB chosen to synthesize the macroinitiator was previously synthesized by Dr. Yaohong Chen using styrene epxoide (SE) according to procedures from Puskas et al. The starting PIB-OH

(sample id: PIB-OH #16) was measured to have Mn = 5,600 g/mol and Mw/Mn = 1.07 by

SEC. Figures 4.4.9 and 4.4.10 shows the 1H and 13C NMR spectra of the starting PIB-

OH, respectively. The doublet of doublet from 3.4~3.6 ppm corresponds to the CH2 (5)

139 next to the primary hydroxyl group. The two peaks (17) at 4.6 and 4.8 ppm corresponds

to olefinic end groups formed by dehydrohalogenation during stroage. The molecular

weight of the PIB-OH was calculated using integration values. The ratio of (9) and (10) is

2:6 and confirmed the IB repeat unit. The integral of hydroyl end group (5) is set as 2.04.

The MW of the polymer calcualted from NMR is ~5300 g/mol, which agrees well with

the values obtained from SEC.

Figure 4.4.9 - 1H NMR spectrum of starting PIB-OH (#16). (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.9).

13 In the C NMR spectrum (Figure 4.4.10), the CH2-OH carbon (5) appears at 71 ppm. The phenyl ring (δ = 124 (1), δ = 129 (2) δ = 126 (3), and δ = 148 (4)) appears at the expected positions. The carbon resonances of the olefinic end group (δ = 27 (16), δ =

113 (17), and δ = 146 (18)) confirms that de-halogenation occurred during storage.

140

13 Figure 4.4.10 – C NMR spectrum of starting PIB-OH (#16). (solvent: CDCl3, nt = 5,000, d1 = 5s, 75 MHz, Appendix: B.4.4.10).

Ms Gautriaud previously attempted to attach the nerol model compound through a bromo-functionalized nerol with low efficiency. She later attempted a tosyl- functionalized compound and showed improved efficiency to ~20% model compound attachment. According to her thesis, the success of this reaction lies in the removal of all traces of water from the reaction.

To remove all traces of water, THF was first distilled from sodium/benzophenone and then cryodistilled with calcium hydride (CaH2). The PN-Ts and PIB-OH were dissolved in small amounts of hexanes and then freeze-dried. All chemicals were stored in air-free glassware and degassed. The reaction proceeded in the glovebox where the moisture level was <1 ppm. The extensive drying of all the reagents yielded ~60%

141 attachment of the PN-OTs to PIB-OH after five days. This was a major improvement

over the previously reported 20%.

Figure 4.4.11 shows the 1H NMR spectrum of PN-PIB after column chromatography. The CH2 (5) of the PIB-OH#16 shifts upfield and becomes a singlet at

3.43 ppm. The CH (j) of the pyran group overlaps with (5); the integral from 3.31~3.65 ppm is 5.48, which is slightly lower than the expect value of 6. The CH2 (h) on the PN-

Ts (δ = 4.14 ppm) shifted upfield to 3.71 ppm has an integral of 1.76.

Figure 4.4.11 - 1H NMR spectrum of PIB-PN (product 5). (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.11).

The 13C NMR spectrum supports the findings from the 1H NMR spectrum. In

Figure 4.4.12, carbon resonances from both PIB-OH #16 and PN are observed. The attachment of PN onto PIB-OH #16 should result in downfield shifts of peaks according 142 to simulation from ACD Chemsketch©. This is observed from the 13C NMR spectrum.

Peak (5) and (H) shifted downfield to 75 and 73 ppm, respectively, compared to previous chemical shifts of 71 and 68 ppm.

Figure 4.4.12 - 13C NMR spectrum of PIB-PN (product 5). (125 MHz, 6,400 scans, d1 = 5s,Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.12).

4.4.5. Deprotection of PIB-PN to PIB-Nerol (Product 6)

Deprotection of the PIB-PN was accomplished by reacting PIB-PN with PPTs

(pyridinium p-toluenesulfonate) in a mixed solvent of isopropylalcohol and hexanes

(2:1). In the original synthetic scheme proposed by Ms Gautriaud, the deprotection should proceed in ethanol. However, the solubiilty of the functionalized polymer was poor in ethanol and the intial deportection efficiency was <10% after 20 hours at room temperature. The deprotection efficiency was raised to ~98% in 24 hours after switching 143 to the mixed solvent system. Figure 4.4.13 shows the 1H NMR spectrum of PIB-Nerol after work up and precipitation of the polymer. The peaks corresponding to the pyran protecting group ((i) and (i)) disappeared. The integral of CH2 on the nerol (h) corresponds closely with integrals of (a) (3.91 ppm, d, 2H, J = 15 Hz), confirming the deprotection. The integral ratio of (h),(a) and (5) is (1.94:2.06:2.03), which confirms that the PIB is functionalized with nerol.

Figure 4.4.13 - 1H NMR spectrum of PIB-Nerol (product 6) (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.13).

Figure 4.4.14 shows the 13C NMR spectrum of PIB-Nerol. The carbon resonances corresponding to the pyran group (δ = 98 (I), δ = 63 (J), and δ = 19 (L) δ = 25 (M), and δ

= 30 (K)) are no longer observed. The deprotection shifts CH2 (A) on the nerol upfield to

144 62 ppm from 65 ppm. The carbon resonances appear at the expected positions,

confirming the structure of the product.

Figure 4.4.14 - 13C NMR spectrum of PIB-Nerol (product 6). (75 MHz, 5,000 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.14).

4.4.6. Synthesis of PIB-Nerol-Br (Product 7)

The functionalization of primary alcohol to pyrophosphate was accomplished by

Chen et al. The authors first converted the primary alcohol to bromine and then phosphorylated the bromine intermediate. A modification of her methodology was adopted for the bromination of PIB-Nerol and the residue was purified by column chromatography (eluent: Hx/EA, 10:1 v/v) on silica gel to yield PIB-Nerol-Br (~0.27g,

60%). The PIB-Nerol was transparent and turned light yellow after the bromination.

Figure 4.4.15 shows the 1H NMR spectrum of PIB-Nerol-Br after column

145 chromatography and re-precipitation. The doublet of doublet from the CH2-OH (a) (δ =

4.1~4.3 ppm) becomes a singlet at 3.9 ppm, corresponding to the protons in the CH2-Br

(a). The integration values of (a), (h) and (5) are (1.90:1.94:2.02), which confirms the expected chemical structure.

Figure 4.4.15 - 1H NMR spectrum of PIB-Nerol-Br (product 7). (300 MHz, 128 scans, d1 = 10s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.15).

13 Figure 4.4.16 shows the C NMR spectrum of PIB-Nerol-Br. The CH2-OH (A)

13 (δ = 62 ppm) shifts upfield to 28 ppm to show the transformation to CH2-Br (A). The C

NMR spectrum confirms conversion of PIB-Nerol to PIB-Nerol-Br.

146

Figure 4.4.16 – 13C NMR spectrum of PIB-Nerol-Br (product 7). (125 MHz, 4,600 scans, d1 = 10s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix:B.4.4.16).

4.4.7. Synthesis of Nerol-PP (Model Reaction for Product 8)

A model reaction was to convert an allylic alcohol to allylic pyrophosphate was

attempted following procedures established by Chen et al. They reacted an allylic

bromine with (Bu4N)3HP2O7 in dry acetonitrile for six hours to attached the

+ + pyrophosphate group with (Bu4N) ion pairs and then convert the (Bu4N) ions to either

+ + H or NH4 using ion exchange column chromatography. The product was then purified

by LC and freeze-dried into a white powder. This model reaction proceeded in the

University of Bordeaux, France. The starting compound, Nerol (a clear oil), was first

converted to Nerol-Br (a clear oil) using Corey-Kim bromination, followed by reacting

147 1 with (Bu4N)3HP2O7 to obtain Nerol-PP (white powder). Figure 4.4.17 shows the H

NMR spectra of Nerol-Br together Nerol-PP. CH2 protons (a) (4.54 ppm, d, 2H) at δ =

3.8 ppm shifted downfield 4.54 ppm. The integrals of CH protons (b) and (f) with CH2-

PP protons (c) are (0.9:0.9:2.20). The downfield shift and the integral values confirmed that Chen et al’s procedures were successful in converting allylic alcohols to allylic pyrophosphates.

Figure 4.4.17 - 1H NMR spectrum of Nerol-PP(model compound of product 8). (300 MHz, 128 scans, d1 = 5s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D).

4.4.8. Synthetic Stretegy to Yield Macroinitiator Using Enzyme Catalysis

Based on discussions with our colleagues at the USDA, it was postulated that longer PIB chain on the macroinitiator may hinder the incorporation of the synthetic PIB-

NPP initiator in in vitro NR biosynthesis. Therefore, synthesis of the macroinitiator with a PIB dimer (2,4,4-trimethyl-pentyl-) head group (Figure 4.4.18, Nerol-PP-DVA-TMP) was proposed. Furthermore, Puskas et al. recently utilized Candida antarctica Lipase B 148 (CALB) to functionalize PIB and PEG. CALB is a commercially available enzyme that is

well characterized and able to facilitate transesterification and Michael addition under diverse conditions. Utilizing enzymatic catalysis in the synthetic scheme would offer an alternative, environmentally-friendly, and efficient pathway to yield the macroinitiator.

Figure 4.4.18 shows the proposed synthetic strategy. After protection of the nerol with the dihydropyran (step 1) and allylic oxidation of PN to yield PN-OH (step 2), PN-

OH was reacted with divinyl adipate (DVA), catalyzed by CALB to yield PN-DVA

(product 9, step 8). PN-DVA was then reacted with TMP-OH (PIB dimer), to yield PN-

DVA-TMP (product 10, step 9). PN-DVA-TMP was deprotected (step 10) to regain the

allylic alcohol functional group (Nerol-DVA-TMP, product 11). Nerol-DVA-TMP was

then brominated (product 12, step 11) and phosphorylated (step 12) to yield the Nerol-

PP-DVA-TMP initiator (product 13). In comparison with the previously proposed

scheme, this alternative method introduced two ester bonds as a linker.

149 O O O O OH O O O divinyl adipapte (DVA) O OO OO CALB (10% wt) 9 Hexanes, 50 oC 3 PN-DVA PN-OH step 8 O O OH O O TMP-OH 12 OO 13 CALB (10% wt) Hexanes, 50 oC PN-DVA-TMP step 9

O O O PPTs O IPA(2):Hx(1) OH step 10 14 Nerol-DVA-TMP

O O O NBS/Me S O 2 Br CH2Cl2 15 step 11 Nerol-Br-DVA-TMP

O (Bu4N)3HP2O7 O O acetonitrile O IXC column PP 16 step 12 Nerol-PP-DVA-TMP

Figure 4.4.18 – Enzyme-catalyzed synthetic scheme to yield Nero-PP-DVA-TMP.

4.4.9. Synthesis of PN-DVA (Product 9)

The first step in synthesizing the Nerol-PP-DVA-TMP is to obtain mono- substituted DVA allylic alcohol of PN (PN-DVA, product 3). Since DVA has two vinyl

functionalities, four molar equivalents of vinyl groups or two times excess of DVA, was

150 reacted with PN-OH to hinder di-substituted PN-DVA-PN formation. The reaction was monitored by TLC and was complete after 24 hours. From the TLC, small amount of

dimer formation was observed. The reaction yielded ~1.3g of PN-DVA, which was ~80%

conversion and ~20% dimer formation. The product was purified by flash column

chromatography (eluent: Hx/THF, 4:1, v/v) (Rf = 0.5 for PN-DVA, Rf = 0.4 for PN-

DVA-PN and Rf = 0.9 unreacted DVA).

Figure 4.4.19 shows the 1H NMR spectrum of PN-DVA. When compared to the

1H NMR spectrum of PN-OH (product 3, Figure 4.4.5), the proton resonances at δ = 3.87

ppm corresponding to the (CH2-OH) protons of PN-OH shifts downfield to δ = 4.46 ppm. Further, the transesterification reaction makes the CH2 protons, (p) and (s) on the

DVA in different environments and becomes two separate triplets at δ = 2.31 and δ =

2.37 ppm, respectively compared to the previous triplet at δ = 2.34 ppm. Proton

resonances attributed to the OCH=CH2 (δ = 4.90 (u)), CH2-OCO (δ = 4.46 (g’)) and O-

CH-O (δ = 4.58 (i)) have integration ratios of (u):(g’+i) as 1.87:4.13.

151

Figure 4.4.19 - 1H NMR spectrum of PN-DVA (product 9). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.19).

13 Figure 4.4.20 shows the C NMR spectrum of PN-DVA. The CH2-O resonance

(D’) connected to DVA in the starting PN-OH shifts downfield from δ = 68 ppm to δ =

72 ppm. New carbon resonances of the vinyl groups (δ = 146 (T), δ = 96 (U)) and

carbonyl carbons resonances of adipic ester (δ = 171 (P’), δ = 173 (S’)) on the DVA appear at the expected positions after 24 hours of reaction. Similarly to the observation

1 from the H NMR spectrum, the carbon resonances of the two CH2 of DVA, (P) and (S),

appears as two separate peaks at δ = 40 (P) and δ = 42 (S) compared to the single peak (δ

= 39 ppm) before the reaction. 13C resonances at the expected positions confirm the

chemical structure of PN-DVA.

152

Figure 4.4.20 – 13C NMR spectrum of PN-DVA (product 9). (125 MHz, 2,800 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.20).

ESI-MS spectrum of PN-DVA (Figure 4.4.21) shows the transesterification products of PN-OH (product 3) with DVA after column chromatography. The calculated

+ monoisotopic mass of PN-DVA is m/z = 431.24 [408.25 (C23H36O6) + 22.99 (Na )]. The formation of a mono-substituted product with m/z = 431.4 [M+Na+] is confirmed.

Another peak at m/z = 221.3 corresponds to unfunctionalized DVA [198.09 (C10H14O4) +

22.99 (Na+)]. There was about 3% of un-substituted DVA and 97% of PN-DVA, which

was within the instrumental error of NMR spectroscopy. The di-substituted PN-DVA-PN

was not observed in ESI-MS and was successfully removed.

153

Figure 4.4.21 – ESI-MS spectrum of PN-DVA (product 9). (solvent: THF, cationizing agent: NaTFA, sample/salt: 100/1 (v/v)).

4.4.10. Synthesis of TMP-OH (PIB dimer)

A (2,4,4-trimethyl-pentyl-) head group was proposed for the macroinitiator and a

PIB dimer (TMP-OH) was synthesized. TMP-OH was synthesized in a three-step synthetic scheme. (Figure 4.3.22) First, TMPCl was synthesized by bubbling HCl gas in bulk. Then, TMPCl converted to TMP-allyl using TICl4 and ATMS in the glove box.

Finally, the TMP-allyl went under hydroboration-oxidation to yield TMP-OH.

154

Figure 4.4.22 – Synthetic scheme to yield TMP-OH (PIB dimer).

The 1H NMR spectrum of the starting compound (TMP-1) is shown in Figure

4.4.23. The vinyl resonances (δ = 4.65, 4.85 (d)) have integral of 2.13. The CH3 protons

(a) and (c) (δ = 0.95 and 1.80 ppm) have a integration ratio of (a):(c) as 9.21:3.20. The hydrogens of the CH2 group (b) show up at δ = 1.96 ppm with integral value of 2.24.

155

Figure 4.4.23 - 1H NMR spectrum of TMP-1. (300 MHz, 32 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D).

TMP-1 was bubbled with HCl gas until 100% conversion to TMPCl (step 1).

TMPCl was neutralized with sodium bicarbonate powder and filtered to yield TMPCl

(~85%, 13.2 g, loss of product from filtering). Figure 4.4.24 shows the 1H NMR spectrum

1 of TMPCl. When compared to the H NMR spectrum of TMP-1 (Figure 4.4.23), CH3 (c) from TMP-1 shifts upfield from 1.80 ppm to 1.67 ppm. The C=CH2 resonances (δ = 4.65,

4.85 (d)) disappear and shift upfield to merge with XC-CH3 (δ = 1.67 (c)) to have a total

integral value of 6.16. The integration ratio of protons (a) and (b) is 9.00:1.93. The 1H resonances appear at the expected positions and integrals, confirming the structure of the product.

156

Figure 4.4.24 - 1H NMR spectrum of TMPCl. (300 MHz, 64 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D).

TMPCl was first reacted with TiCl4 under the presence of DtBP (proton trap), and then terminated with ATMS to yield TMP-allyl (step 2). After washing and work-up, the

residue was purified by flash chromatography (eluent: diethyl ether/hexane, 1:9 v/v;

TLC: Rf = 0.6 for TMP-allyl and Rf = 0.3 for ATMS) on alumina to yield TMP-allyl

(~90%, 4.12 g). Figure 4.4.25 shows the 1H NMR spectrum of TMP-allyl. In 1H NMR spectrum, new proton resonances at δ = 4.92 and δ = 5.76 ppm corresponding to the

CH=CH2 (e) and CH=CH2 (f) appear with integration ratio of (f):(e) as 2.13:0.99. The

CH3 (c) resonances shift upfield from 1.67 ppm to 1.36 ppm. No peaks were observed near 0 ppm to confirm that the excess ATMS was successfully removed from the residue.

157

Figure 4.4.25 - 1H NMR spectrum of TMP-allyl. (300 MHz, 32 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D).

TMP-allyl went under hydroboration-oxidation with 9-BBN and H2O2 to yield

TMP-OH (step 3). The residue was purified by flash chromatography (eluent: ethyl acetate/hexane, 1:6 v/v; TLC: Rf = 0.3 for TMP-OH, Rf = 0.7 for TMP-allyl, and Rf = 0.4 for TMP-9-BBN intermediate) on alumina to yield TMP-OH (~72%, 3.12 g). In 1H NMR spectrum (Figure 4.4.26), new proton resonances at δ = 4.27 and δ = 3.38 ppm corresponding to the CH2-OH (g) and CH2-OH (f) appear with integration ratio of (f):(e)

as 1.01:2.00. The vinyl 1H resonances (δ = 4.92 (f) and δ = 5.76 ppm (e)) from TMP-allyl

(Figure 4.4.25) disappeared. The CH=CH2 (e) from TMP-allyl shifted upfield to δ = 1.41

ppm after oxidation. The integration ratio of (a):(b):(c):(e) is 8.96:2.19:6.30:2.05. Proton

158 resonances at the expected positions and integral values confirm the chemical structure of

TMP-OH for the synthesis of PN-DVA-TMP.

Figure 4.4.26 - 1H NMR spectrum of TMP-OH. (300 MHz, 32 scans, d1 = 1s, Pulse angle = 45o, T = 25°C, Solvent: chloroform-D).

The ESI-MS spectrum of TMP-OH is shown in Figure 4.4.27 after column chromatography. The calculated monoisotopic mass of TMP-OH is m/z = 195.17 [172.18

+ (C11H24O) + 22.99 (Na )]. The successful synthesis of TMP-OH with m/z = 195.6

[M+Na+] is confirmed. Another peak at m/z = 267.3 corresponds to boron impurities (9-

BBN, [244.03 (C16H30B2) + 22.99 (Na+)] = 267.02) from the hydroboration reaction.

159

Figure - 4.4.27 ESI-MS of TMP-OH. (solvent: Methanol, cationizing agent: NaTFA, sample/salt: 100/1 (v/v)).

4.4.11. Synthesis of PN-DVA-TMP (Product 13)

The second step in synthesizing the Nerol-PP-DVA-TMP is to attach TMP-OH

to PN-DVA (product 9) to yield PN-DVA-TMP (product 13). Since PN-DVA only has

one vinyl group for transesterification, a 1.1 molar equivalent of PN-DVA (2.03 g, 0.5

mmol) to TMP-OH (product 12) (0.78 g, 0.46 mmol) was used. The reaction was

performed at 50 oC in THF under nitrogen atmosphere. The reaction was monitored by

TLC and was complete after 5 hours. CALB was filtered using a 0.45 µm PTFE syringe filter and then concentrated by rotary evaporation. The reaction yielded ~1.84 g (3.45 mmol) of PN-DVA-TMP, which was ~70% conversion. On the TLC, a very small spot of excess PN-DVA was observed. The product was purified by silica column

chromatography (eluent: Hexane/THF, 4:1, v/v) (Rf = 0.5 for PN-DVA, Rf = 0.3 for PN-

DVA-TMP). 160 Figure 4.4.28 shows the 1H NMR spectrum of PN-DVA-TMP. New proton resonances at δ = 4.08 appears and corresponds to the CH2-O (1) between TMP-OH and

DVA. The integration ratio of (a) (4.21 ppm), (j), (3.52 ppm) and (1) (4.08 ppm) should be 2:2:2. Since there is overlapping between the peaks, the sum of the integrals is 6.08

(3.87 + 2.21), which is close to the expected value. Three new peaks attributed to the

TMP-OH (δ = 0.99 (7), δ = 1.12 (4’), and δ = 1.42 (5)) are observed. The ratio of integrals between (7) and (4’) is 8.78:5.87, which confirms the desired tetramethyl hexyl head group. The vinyl end group resonances initially present on the PN-DVA (δ = 4.90

(u) and δ = 7.28 ppm (t)) are not observed. The CH2 protons, (p) and (s), corresponds to two overlapping triplet peaks at δ = 2.29 and δ = 2.37 ppm, respectively, due to the different environments in which they are located.

Figure 4.4.28 - 1H NMR spectrum of PN-DVA-TMP (product 13). (300 MHz, 32 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.28). 161 Figure 4.4.29 shows the 13C NMR spectrum of PN-DVA-TMP. The vinyl carbon resonances (δ = 146 (T), δ = 96 (U)) from PN-DVA (Figure 4.3.20) disappeared. New peaks attributing to the tetramethyl hexyl head group (δ = 28 (5), δ = 31 (3), δ = 18 (5’), δ

= 22 (3’), and δ = 51 ppm (4)) are found. The carbon on the TMP-OH next to the hydroxyl group CH2-O (δ = 66 ppm (1)) is also observed. The CH2-CH2-C (2) on the

TMP-OH appears at δ = 37 ppm. One of the carbonyl carbons resonances of adipic ester

from PN-DVA (δ = 173 (S’)) shifts slightly upfield to 172 ppm. 13C resonances at the expected positions confirm the chemical structure of PN-DVA.

Figure 4.4.29 – 13C NMR spectrum of PN-DVA-TMP (product 13). (125 MHz, 3600 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.29).

Figure 4.4.30 shows the ESI-MS spectrum of PN-DVA-TMP after column chromatography. The calculated monoisotopic mass of PN-DVA-TMP is m/z = 559.40

+ [536.41 (C32H56O6) + 22.99 (Na )]. The formation of PN-DVA with a tetramethyl hexyl head group is confirmed with a major peak at m/z = 559.13 [M+Na+]. Another peak at

162 + m/z = 575.14 corresponds to the potassium complex [536.41 (C32H56O4) + 38.96 (K ) =

575.37].

Figure 4.4.30 – ESI-MS spectrum of PN-DVA-TMP (product 13). (solvent: THF, cationizing agent: NaTFA, sample/salt: 100/1 (v/v)).

4.4.12. Synthesis of Nerol-DVA-TMP (Product 14)

The deprotection of PN-DVA-TMP to Nerol-DVA-TMP proceeded in a mixed solvent of isopropyl alcohol(IPA) and hexane (IPA:Hexanes, 2/1, v/v). The original scheme was to deprotect the PN-DVA-TMP in ethanol; however, the product could not becompletely dissolved in ethanol. PN-DVA-TMP (1.05g, 2.0 mmol) was first dissolved in mixed solvent of IPA:Hexanes (10.0 mL). After 10 minutes of stirring, PPTs (0.2 g,

0.8 mmol) dissolved in 5.0 mL IPA:Hexanes were injected through a rubber septum at room temperature and left stirring for 20 hours. The reaction was monitored by TLC and was complete after 20 hours. The reaction yielded ~0.8 g (1.8 mmol) of Nerol-DVA-

163 TMP, which corresponded to ~90% conversion. On the TLC, other than the product

peak, a spot corresponding to 1-hydroxy pyran from the acidic ring opening of the pyran

group during deprotection) was observed. The product was purified by column chromatography (eluent: Hexane/THF, 4:1, v/v) (Rf = 0.1 for Nerol-DVA-TMP, Rf =

0.35 for 1-hydroxy pyran).

Figure 4.4.31 shows the 1H NMR spectrum of Nerol-DVA-TMP. Proton

resonances at δ = 4.85 and 3.52 corresponding to the O-CH-O (i) and O-CH2-CH2 (j) of

the pyran protecting group of PN-DVA-TMP disappeared. The same is observed for

peaks at δ = 1.83 and 1.54 from the pyran ring. The integration ratio of (b+f), (h), (1) and

(a) (3.81 ppm, d, 2H) is 2:2:2:2, which confirms the structure of Nerol-DVA-TMP. The

O-CH2-CH2 (1) (4.08 ppm, t, 2H) splits into triplets due to two adjacent protons (2) and

CH-CH2-OH (a) splits into due to adjacent protons on (b). The desired tetramethyl hexyl

head group has integrals of (7) and (4’) as 9:6 in relation to (b+f), which confirms that

the TMP was not cleaved during the pyran deprotection process. The CH2 protons, (o)

and (r), appear as two overlapping triplet peaks at δ = 2.29 and δ = 2.37 ppm, respectively, due to the different environments in which they are located.

164

Figure 4.4.31 - 1H NMR spectrum of Nerol-DVA-TMP (product 14). (300 MHz, 64 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix B: 4.4.31).

Figure 4.4.32 shows the 13C NMR spectrum of Nerol-DVA-TMP. The carbon resonances (δ = 108 (I), δ = 63 (J), δ = 27 (K), δ = 20 (L), and δ =31 (M)) from pyran group in PN-DVA-TMP (Figure 4.3.28) disappeared. The carbons attributing to the tetramethyl hexyl head group (δ = 28 (7), δ = 31 (4’), δ = 18 (6), δ = 22 (4), and δ = 51 ppm (5)) are intact. The CH-CH2-OH (A) on the nerol shifted downfield to δ = 67 ppm.

13C resonances at the expected positions confirms the chemical structure of Nerol-DVA-

TMP.

165

Figure 4.4.32 – 13C NMR spectrum of Nerol-DVA-TMP (product 14). (75 MHz, 3600 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.32).

4.4.13. Synthesis of Nerol-Br-DVA-TMP (Product 15)

The bromination of Nerol-DVA-TMP to Nerol-Br-DVA-TMP proceeded in

dichloromethane at -40 oC using N-bromosuccinimide (NBS) and dimethyl sulfide.

Nerol-DVA-TMP (0.55 g, 1.2 mmol) and NBS (0.27 g, 1.5 mmol) were added to CH2Cl2 and left stirring for 20 minutes before warming the reaction flask to 0 oC. After the

o temperature stabilized at 0 C, dimethyl sulfide (Me2S, 0.1 g, 1.6 mmol) was injected and the reaction proceeded for 6 hours. The reaction yielded ~0.4 g (0.8 mmol) of Nerol-Br-

DVA-TMP, which corresponded to ~65% conversion. The product was purified by column chromatography (eluent: Hexane/Ethyl acetate (EA), 10:1, v/v) (Rf = 0.25 for

Nerol-Br-DVA-TMP, Rf = 0.1 for Nerol-DVA-TMP).

Figure 4.4.33 shows the 1H NMR spectrum of Nerol-Br-DVA-TMP. The proton resonance δ = 3.81 ppm (–CH2-OH) of Nerol-DVA-TMP shifted upfield to 3.54 ppm to

166 signify the conversion to –CH2-Br. The integration ratios of (b+f), (h), (1) and (a) remain

2:2:2:2, which confirms the structure of Nerol-Br-DVA-TMP. The CH2-Br (a) (3.54

ppm) splits into a triplet from the adjacent proton on (b). The desired tetramethyl hexyl

head group retained integrals of (7) and (4’) as 9:6 in relation to (b+f) on the Nerol-Br-

DVA-TMP, which confirms that it was not cleaved during the deprotection and

bromination process.

Figure 4.4.33 - 1H NMR spectrum of Nerol-Br-DVA-TMP (product 15). (300 MHz, 64 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix B: 4.4.33).

Figure 4.4.34 shows the 13C NMR spectrum of Nerol-Br-DVA-TMP. The carbon resonances next to the alcohol (δ = 67 ppm (A), (Figure 4.3.31)) in Nerol-DVA-TMP

disappeared. A new 13C resonance appeared at δ = 67 ppm (A), confirming the

13 13 transformation from CH-CH2-OH to CH-CH2-Br. The C resonances matched the C

167 NMR spectrum of Nerol-DVA-TMP at the expected positions confirms the chemical

structure of Nerol-Br-DVA-TMP.

Figure 4.4.34 – 13C NMR spectrum of Nerol-Br-DVA-TMP (product 15). (75 MHz, 3600 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix B: 4.4.34).

4.4.14. Synthesis of Nerol-PP-DVA-TMP (Product 13) (Chen’s Method150)

The phosphorylation of Nerol-Br-DVA-TMP to Nerol-OPP-DVA-TMP proceeded in air-free conditions for 6 hours, which was in accordance with literature.150

Nerol-Br-DVA-TMP (0.3 g, 0.6 mmol) and (Bu4N)3HP2O7 (0.9 g, 1 mmol) was added to

31 column (IXC) (NH4+ form). Figure 4.4.35 presents the P NMR spectrum of the crude product in CDCl3. Four phosphorous peaks were observed; peaks at δ = -0.64 (X) and δ =

168 -10.15 (Y) ppm correspond to the starting (Bu4N)3HP2O7. The diphosphate synthetic initiator intermediate had two 31P peaks at δ = -11.04 (X’) and δ = -12.17 (Y’) ppm. The

tetrabutylammonium counter cations needed to be exchanged to a hydrogen counter ion

to obtain the pyrophosphate end group. However, after passing through the IXC column,

crude product of Nerol-PP-DVA-TMP was found to be disintegrated or decomposed

under the acidic conditions. The ester bonds present on the DVA showed acid catalyzed

hydrolysis from HBr from the byproduct of Nerol-Br-DVA-TMP and (Bu4N)3HP2O7. It is also plausible that the acid groups on the DOWEX column material were responsible

for the cleavage. Therefore, an alternative method of phosphorylation was required to

obtain the diphosphate initiator.

Figure 4.4.35 – 31PNMR spectrum of diethyl chlorophosphate. Bu denotes n-butyl group. (121.5 MHz, 300 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

169 4.4.15. Synthetic scheme to yield Nerol-OPP-DVA-TMP (Product 16) (Coates’

Method151)

Since the ester bonds of the macroinitiator are prone to hydrolysis under acidic

conditions during the ion exchange step. In addition, the bromine end group on the Nerol-

Br-DVA-TMP also contributed to the disintegration as an acid generation source.

Therefore, to obtain the synthetic initiator, another synthetic route was attempted. The

revised chemical scheme to yield the diphosphate end group follows steps previously

developed by Coates et al. in which a primary phosphate intermediate was used instead of the Br.151 (Figure 4.4.36) Nerol-DVA-TMP (Product 11) is first reacted with diethyl

chlorophosphate to yield Nerol-OP(O)(OEt)2-DVA-TMP (Product 14). Then, Product

14 is reacted with (Bu4N)3HP2O7 in anhydrous acetonitrile to yield the macroinitiator,

Nerol-OPP-DVA-TMP (Product 13). Using this method, the reactions are in milder

conditions and thus reduce the chances that the ester bonds would cleave by hydrolysis.

170

Figure 4.4.36 – Synthetic scheme to yield Nero-PP-DVA-TMP adapted from Coates et al.

4.4.16. Synthesis of Nerol-OP(O)(OEt)2 (Model Reaction)

Model reactions were employed to test the feasibility of the proposed synthetic scheme shown in Figure 4.4.36. The model compound used was nerol because it is structurally similar to the macroinitiator and readily available. A solution of Nerol (250 mg, 1.62 mmol) and pyridine (0.16g, 2.02 mmol) in chloroform (5 mL) was solubilized at

o 1 C. Then, diethyl chlorophosphate solution (0.28 g, 1.62 mmol in 1 mL CHCl3) was added slowly and stirred for 4 hours. The product was purified by silica gel column (1:3

171 Hexane/Ethyl acetate, Nerol-P(O)(OEt)2, Rf = 0.34). Nerol-P(O)(OEt)2was a light yellow

Figure 4.4.37 shows the 31P NMR of diethyl chlorophosphate, which is the starting

reagent. Phosphorous resonances at δ = 4.38 ppm corresponds to the phosphorous atom

(V). After the reaction proceeded for four hours at 1 oC, the 31P NMR spectrum of Nerol-

31 OP(O)(OEt)2 was taken pre-purification to observe the P peaks of the crude product, as shown in Figure 4.4.37.

Figure 4.4.37 – 31PNMR spectrum of diethyl chlorophosphate. (121.5 MHz, 400 scans, d1 = 2s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

From Figure 4.4.38, two phosphorous peaks were observed, δ = 4.39 and δ = -

13.18 ppm. δ = -13.18 ppm should correspond to the nerol primary phosphate. When

Coates et al. reacted their diol compound with diethyl chlorophosphate, they observed the

product 31P peak to be δ = -6.32 ppm.

172

31 Figure 4.4.38 – PNMR spectrum of crude product of Nerol-P(O)(OEt)2. (121.5 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

Figure 4.4.39 presents the 31P NMR spectrum of the product after purification by column chromatography in alumina with acetonitrile. Only one phosphorous peak was observed at δ = -13.18 ppm. Figure 4.4.40 shows the 1H NMR spectrum of Nerol-

1 P(O)(OEt)2. The H NMR spectrum of the nerol starting compound can be seen in

Figure 4.4.2.

173

31 Figure 4.4.39 – PNMR spectrum of Nerol-P(O)(OEt)2. (after column) (121.5 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

1 Figure 4.4.40 show the H NMR spectrum of Nerol-P(O)(OEt)2. Proton resonances at δ = 5.35 and 3.13 ppm corresponding to the two CHs (b) (5.42 ppm, t, H) and (f) (5.08 ppm, t, H) of the nerol. The CH2-OH (a) (4.43 ppm, d, 2H) of Nerol shifted

downfield from δ = 4.11 ppm to δ = 4.46 ppm in Nerol-P(O)(OEt)2, which confirmed the primary phosphate end group on the nerol. New 1H resonances at δ = 3.82 and δ =

1.00 ppm corresponds to the O-CH2-CH3 (t) and O-CH2-CH3 (u) (1.01 ppm, t, 3H) on the

diethyl phosphate end group. The O-CH2-CH3 (t) (3.86 ppm, m, 2H) splits into multiplets

due to three adjacent protons (u) and O-CH2-CH3 (u) splits into a triplet from adjacent

two protons on (t). The integration ratio of (b), (a), (t) and (u) is 1:2:4:6, which confirms the structure of Nerol-P(O)(OEt)2.

174

1 Figure 4.4.40 - H NMR spectrum of Nerol-P(O)(OEt)2. (300 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.40).

4.4.17. Synthesis of Nerol-PP (Model Reaction)

Nerol-P(O)(OEt)2 was reacted with (Bu4N)3HP2O7 in anhydrous acetonitrile for 5 hours to obtain Nerol-PP. Figure 4.4.41 shows the 31P NMR spectrum of the crude

31 product. The starting reagent ((Bu4N)3HP2O7) has P NMR peaks at δ = -0.64 (X) and -

10.15 (Y) ppm. Phosphorous NMR peaks appeared at δ = -11.04 (X’) and -12.17 (Y’) ppm respectively which are assigned to nerol diphosphate (Nerol-PP). However, a new phosphorus peak was also detected from the crude product at 2.13 ppm (W’). According to the literature, this peak corresponded to the monophosphate alcohol given off during the reaction. The desired Nerol-PP compound was isolated by silica gel column chromatography (eluent: anhydrous acetonitrile) (Rf = 0.55 for Nerol-PP, Rf = 0.9 for 175 31 Nerol- P(O)(OEt)2 ). The observed peaks in the P NMR spectrum of the Nerol-PP were (X’), (Y’) and a trace amount of (W). (not shown)

Figure 4.4.41 – 31PNMR spectrum of Nerol-PP. (crude, Bu denotes n-Butyl). (121.5 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

Figure 4.4.42 shows the ESI-MS spectrum of Nerol-PP after column chromatography in the negative mode. The solvent was anhydrous acetonitrile and no ionizing agent was used because the compound was a negatively charged entity with tetra-n-butylammonium counter cations. Initially, the ESI-MS spectrum only showed a single peak at m/z = 241.7 Da, which corresponded to the tetra-n-butylammonium cation.

(C16H36N = 242.28 Da). This was probably because the tetra-n-butylammonium cations flooded the detector. The ESI mass spectrum in negative mode shows only a

- C10H17O7P2 ion (m/z = 311.0), consistent with the nerol diphosphate structure. The actual 176 3- anion in the sample analyzed is C10H17O7P2 , indicating that the partial oxidation to

- C10H17O7P2 take place during the ESI process.

Figure 4.4.42 – ESI-MS spectrum of Nerol-PP. (Negative Mode. solvent: acetonitrile, ionizing agent: n/a).

4.4.18. Synthesis of Nerol-OP(O)(OEt)2-DVA-TMP (Product 15)

After confirming that the model reaction could successfully attach a diphosphate onto the Nerol-PP, phosphorylation of Nerol-DVA-TMP was carried out. At first, Nerol-

DVA-TMP was reacted with diethyl chlorophosphate in dichloromethane at room

temperature with pyridine as the acid scavenger. The reaction proceeded at 1 oC for 4 hours and was monitored by 31P NMR. It was observed that the signal at δ = 4.37 ppm attributed to diethyl chlorophosphate shifted up field to δ = -12.76 ppm in the product

Nerol-OP(O)(OEt)2-DVA-TMP. It was observed that after 1 ½ hour of the reaction, the relative intensities of the 31P NMR peaks did not change, which suggested that the

177 reaction stopped in around 1 ½ hour. The reaction yielded ~150 mg (~50% yield) of

Nerol- OP(O)(OEt)2-DVA-TMP after purification by silica gel flash chromatography

(eluent: dichloromethane, Rf = 0.6 for Nerol-OP(O)(OEt)2-DVA-TMP, Rf = 0.25 for

Nerol-DVA-TMP.)

Figure 4.4.43 shows the 31PNMR spectrum of Nerol-OP(O)(OEt)-DVA-TMP.

The peak at δ = 4.37 ppm (V) is from the excess starting material, diethyl

chlorophosphate. The peak at δ = -12.76 (W) ppm corresponds to the attached

monophosphate end group in Nerol-OP(O)(OEt)-DVA-TMP.

31 Figure 4.4.43 – PNMR spectrum of Nerol-OP(O)(OEt)2-DVA-TMP. (121.5 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

1 The Figure 4.4.44 show the H NMR spectrum of Nerol-P(O)(OEt)2-DVA-TMP.

peak at δ = 5.42 ppm corresponds to the two CHs protons (b) and (f) of Nerol-

P(O)(OEt)2-DVA-TMP . The CH2-O-PP (a) shifted downfield from δ = 3.81 ppm in

178 Nerol-DVA-TMP to δ = 4.88 ppm, which confirmed the primary phosphate end group

on the Nerol-P(O)(OEt)2-DVA-TMP . New peaks at δ = 3.74 and δ = 1.06 ppm correspond to the O-CH2-CH3 (t) and O-CH2-CH3 (u) protons respectively on the diethyl

phosphate end group. The O-CH2-CH3 (t) split into multiplets due to the three adjacent

protons on O-CH2-CH3 (u) and (u) in turn split into triplets from two adjacent protons on

(t). The integration ratio of (b+f), (a), and (t) was found to be 2:2:4, which confirmed the structure of Nerol-P(O)(OEt)2-DVA-TMP. Since the compound was not stable and easily go under hydrolysis, 13C NMR and ESI-MS of the compound was not taken before proceeding with the next step of the synthesis.

1 Figure 4.4.44 - H NMR spectrum of Nerol-P(O)(OEt)2-DVA-TMP. (300 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.44).

179 4.4.19. Synthesis of Nerol-PP-DVA-TMP (Product 16)

The final step of the synthetic initiator synthesis involved reacting the purified

Nerol-P(O)(OEt)2-DVA-TMP with (Bu4N)3HP2O7 in anhydrous acetonitrile. A detailed

procedure of the reaction is described in Chpater 3.3.16. The previously proposed and

discussed bromine intermediate step (Figure 4.4.18 in Chapter 4.4.8) and use of ion

exchange column to exchange tetra-n-butylammonium cations with protons was not used

in order to avoid any hydrolysis of the DVA ester bonds of the product Nerol-PP-DVA-

TMP. The reaction proceeded at room temperature for 2 days and was monitored by 31P

NMR under air-free conditions with molecular sieves to remove water. The reaction was

31 monitored by relative P NMR peak intensities from the starting material, (Bu4N)3HP2O7

(δ = -0.64 (X) and -13.17 ppm (Y)) to peaks at δ = -11.04 (X’) and -12.17 (Y’) ppm of

the product Nerol-P(O)(OEt)2-DVA-TMP. The reaction yielded ~240 mg (~54% yield) of Nerol-PP-DVA-TMP, after purification by cellulose chromatography. (Whatman CF-

11, eluent: anhydrous acetonitrile, Rf = 0.4 for Nerol-OP(O)(OEt)-DVA-TMP, Rf =

0.15 for Nerol-PP-DVA-TMP.) From the 31P NMR spectrum, the reaction showed ~90% conversion into the product Nerol-PP-DVA-TMP; however, only ~54% of it could be recovered after column chromatography, it appeared that the product Nerol-

OP(O)(OEt)-DVA-TMP had a good affinity to cellulose and could not be entirely

removed from the cellulose column under the current applied conditions.

Figure 4.4.45 shows the 31P NMR spectrum of Nerol-PP-DVA-TMP after cellulose chromatogrphy. The resonance at δ = -11.04 (X’) and -12.17 (Y’) ppm corresponded to the attached pyrophosphate end group on the Nerol-PP-DVA-TMP. The

180 peak at δ = 2.12 (W) corresponded to the diethylhydroxyphophsate that was given off as

the leaving group when Nerol-P(O)(OEt)2-DVA-TMP was reacted with (Bu4N)3HP2O7.

Figure 4.4.45 – 31PNMR spectrum of Nerol-PP-DVA-TMP. (product 16). (121.5 MHz, 400 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D).

Figure 4.4.45 show the 1H NMR spectrum of Nerol-PP-DVA-TMP. The peak at

δ = 4.88 ppm for CH2-O-PP (a) (Figure 4.4.45) shifted slightly upfield compared to

Nerol-PP-DVA-TMP (4.79 ppm) and the integration of (a) was 2.09 relative to the integral of 2.00 for (b+f). The peaks at δ = 3.74 and δ = 1.06 ppm corresponding to the

O-CH2-CH3 (t) and O-CH2-CH3 (u) Nerol-OP(O)(OEt)2-DVA-TMP (Figure 4.4.43)

disappeared and two new peaks were observed at δ = 3.32 (w) and 0.93 (z) ppm respectively in Figure 4.4.45. These two peaks corresponded to the N-CH2-CH3-CH2-CH3

181 protons on the tetra-n-butylammonium counter cation of Nerol-PP-DVA-TMP with

integral values of 24.16 and 45.08 respectively. The (x) and the (y) protons overlaps with the aliphatic protons on the TMP head group and were observed as increase in relative integrals on the 1H NMR spectrum from 16.13 to 65.02. The other remaining protons in the 1H NMR spectrum (Figure 4.4.45) matched well with the expected positions and integration values for Nerol-PP-DVA-TMP thereby confirming the chemical structure and successful synthesis of the desired product, Nerol-PP-DVA-TMP.

Figure 4.4.46 - 1H NMR spectrum of Nerol-PP-DVA-TMP. (product 16). (300 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.46).

Figure 4.4.47 shows the 13C NMR spectrum of Nerol-PP-DVA-TMP. The carbon

resonances next to the diphosphate was observed at δ = 62 ppm (A). 13C resonances appeared at δ = 60 (W), 28 (X), 22 (Y), and 13 (Z) ppm confirming the presence of tetra- 182 n-butylammonium ion in the product and the chemical structure of Nerol-PP-DVA-

TMP.

Figure 4.4.47 – 13C NMR spectrum of Nerol-PP-DVA-TMP (product 16). (75 MHz, 64 scans, d1 = 1s, Pulse angle = 90o, T = 25°C, Solvent: chloroform-D, Appendix: B.4.4.47).

Figure 4.4.48 shows the ESI-MS spectrum of Nerol-PP-DVA-TMP after

cellulose chromatography. The solvent was anhydrous acetonitrile and no cationizing

agent was used. Similarly to the model reaction, the ESI-MS spectrum only showed a

single peak at m/z = 241.7 Da (tetra-n-butylammonium cation, C16H36N = 242.28 Da) when no detection limit was set (Figure A.2). Therefore, to resolve the problem, the ESI mass spectroscopy was setup in a way that only peaks having m/z values between

500~3000 Da were recorded. The peak at m/z = 1336.2 Da corresponded closely to the calculated monoisotopic mass of Nerol-PP-DVA-TMP (m/z = 1336.2 Da [(Mass of

183 3- + + + C27H47O11P2 = 609.26) + 3(Mass of Bu4N ) + H , Mass of Bu4N = 242.28]). For

multiple-charged molecules, m/z is analyzed as “m/z= (MW+nI)/n”, where “MW” represents the molecular weight of the central multiple charged compound, “n” represents the number of stabilizing counter ions and “I” represents the mass of the counter ion. The peak at m/z = 547.7 Da was found to be a doubly charged Nerol-PP-DVA-TMP as the peaks showed an isotope spacing of 0.5 Da. (m/z = 547.7 [m/z = (MW + nI)/n, where n =

3- 2 and I = 242.28. Thus, MW = 610.8], Nerol-PP-DVA-TMP (C27H47O11P2 ) has a monoisotopic mass of 609.26.) Similarly to the model compound (Figure 4.4.41), peaks corresponding to the loss of one tetra-n-butylammonium ion were found. The peak at m/z

= 1095.1 Da corresponded to singly charged Nerol-PP-DVA-TMP but with a loss of one

+ 3- + Bu4N unit (MW = 242.28 Da) and two protons (C27H47O11P2 + 2Bu4N + 2H). The loss

+ of two Bu4N units for Nerol-PP-DVA-TMP were also observed at m/z = 853.4 Da. (m/z

3- + + = 853.4 [853.4 = 609.63(C27H47O11P2 ) + 242.28(Bu4N ) + 2.02(H ) = 853.93).

Interestingly, the peak at m/z = 612.72 corresponded to the synthetic initiator, Nerol-

PP_DVA-TMP where all the tetra-n-butylammonium counter ions are replaced by

3- + protons [609.63Da (Mass of C27H47O7P2 ) + 3.03 Da (Mass of 3H ) = 612.64 Da].

Overall, the ESI mass spectrum in positive mode shows the M+ ion of Nerol-PP-

DVA-TMP (observed at m/z 1336.2; calcd. 1336.11), formed by oxidation of one

+ phosphate anion during the ESI process. Cations missing 1-3 Bu4N counterions are also

+ + observed. All these ions are 1 charge, consistent with replacement of Bu4N (242 Da) by

H+ (1 Da) and phosphate oxidation.

184

+ Figure 4.4.48– ESI-MS spectrum of Nerol-PP-DVA-TMP. Bu4N denotes tetra-n- butylammonium ion. (solvent: acetonitrile, cationizing agent: n/a).

Thus, the detailed spectroscopic analysis of the 1H NMR (Figure 4.4.46) and 13C

NMR (Figure 4.4.47) spectra and ESI-MS (Figure 4.4.48) spectrum as discussed above confirmed that the synthetic initiator Nerol-PP-DVA-TMP was successfully synthesized.

185 CHAPTER V

CONCLUSIONS

The objective of this research was to revisit classical biochemical mechanism of

NR biosynthesis and use modern chemical techniques, such as in vitro NR biosynthesis, to further the understanding of the mechanism that had been elusive to the polymer chemistry community. Puskas et al. proposed that the biosynthesis of NR proceeds by a natural living carbocationic polymerization mechanism (NLCP), in which the dormant pyrophosphate chain ends are reversibly activated by the rubber transferase enzyme and

divalent cation cofactors. This thesis studied the NR biosynthesis in two ways: 1)

addition of IP and other derivatives to shift the enzymatic equilibrium, and 2)

introduction of a synthetic initiator to the in vitro NR biosynthesis system to yield a PIB-

b-NR diblock copolymer.

For the first portion of this thesis, NRs synthesized in vitro were analyzed by high

resolution SEC. Washed rubber particles from RRIM600 were shown to have ~33% gel

content. The soluble rubber had approximately 50% high MW fraction between 105 and

3x106 g/mol, and 50% low molecular weight components with MW between 400 and

4,000 g/mol. In the presence of both FPP initiator and IPP monomer, NR growth was

observed by high-resolution SEC which was able to detect growth of both high and low

MW rubber.

186 Rubber growth was also supported by mass balance measurements, showing

31.6% mass gain. The high-resolution SEC provided new insight into NR biosynthesis,

and will be an efficient characterization technique for future studies. MALDI-ToF MS

was utilized to study the low MW material in NR; it was found that the low MW material

consists of oligoisoprenoids.

In the terpenoid biosynthesis lifecycle, when excess IPP is present, the

equilibrium shifts toward DMAPP which in turn is converted into IP by the isoprene

synthase enzyme. IP is an ideal compound to remove excess IPP via the shifts in the equilibria for its ease of evaporation (boiling point of IP = 34 oC). Puskas et al.

hypothesized that IPP could be generated in situ by “flooding” the active NR producing

latex with IP. This proposal was studied by HR-SEC, gravimetric analysis, in situ Raman spectroscopy and NMR spectroscopy. When IP was introduced to NR latex and WRP, a new peak in ~105 g/mol region was observed in many experiments. In many instances,

mass gains were observed after the addition of IP. When the reaction was monitored by in

situ Raman spectroscopy, we observed decrease in the IP signal and an increase in the

PIP signal. By NMR spectroscopy, only cis-1,4 PIP was observed. Further, when control

experiments with Amylene were implemented, no mass gain or changes in the Raman

signal were observed. Experiments using deuterated IP (D-IP) was conducted to prove

the incorporation of IP into NR. However, we did not observe any trace of D-IP despite

observing mass gain. The results were very promising for future studies and we are

planning to study the experiment with 13C IP to explore the fundamental mechanisms of

terpenoid biosynthesis.

187 The second goal of this research was to design and synthesize a synthetic initiator

for the future synthesis in vitro of a novel polyisobutylene-block-cis-1,4-polyisoprene

(PIB-b-NR) diblock copolymer. First, PIB-OH samples made by “traditional” living

carbocationic polymerization using a -methylstyrene epoxide (MSE)/TiCl4 system to

initiate isobutylene polymerization were used and a PIB-Nerol product was successful

synthesized. However, after much discussion enzymatic synthetic pathway using CALB

was employed to yield the final synthetic initiator with a TMP (PIB-dimer) head group.

The synthetic initiator with functionalized with TMP head group and a model NR end

group (allylic pyrophosphate) was successfully synthesized in a 7-step synthetic scheme.

In the future, this synthetic initiator would be used in in vitro NR biosynthesis to prove the concept to yield a PIB-b-NR diblock copolymer.

188

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201

APPENDIX

Figure A.1 – 1H and 13C NMR chemical shifts from the literature cis 1,4-polyisoprene

1H, Hevea rubber, 25oC, benzene, 5% w/w solution, 800 scans, d1=10sec.152

13C, Hevea rubber, 50oC, benzene, 30% w/w solution, 1,000 scans, d1=2sec.153 ------trans 1,4-polyisoprene

Balata rubber, 25oC, benzene, 5% w/w solution, 800 scans, d1=10sec.153

13C, Gutta percha, 50oC, benzene, 30% w/w solution, 1000 scans, d1=2sec.153 ------

202 3,4-polyisoprene

1H, Anionic synthetic PIP, 25oC, benzene, 40 mg/mL solution, d1=2sec.154

13 C, Synthetic PIP using iron-based catalyst, C2D2Cl4, 10 mg/mL solution *note: high temperature NMR, the authors did not provide more information.155 ------

1,2-polyisoprene

1H and 13C, Anionic synthetic PIP, no information on conditions.156 ------

203

Figure A.2 – ESI-MS of Nerol-PP-TMP-DVA (no data cut-off) (Solvent: acetonitrile, cationizing agent: n/a)

204 Table A.1 - Summary of types of rubber latex components used in in vitro NR biosynthesis systems

Type of Natural Rubber Description Crepe31 Coagulated NR by treating with sodium hydrogen sulfite. Then, the coagulated NR is washed with water. This method removes the majority amounts of the serum constituents which can cause rotting. High-ammonia latex31 Preserved with 0.7% of ammonia on latex phase. Dry rubber content >60% Deproteinized HA latex31,80 Treatment of HA latex with 0.04% (w/v) Alcalase 2.0T (A serine endopeptidase isolated from Bacillus subtilis to remove proteins) and Triton X-100 (nonionic surfactant) at 37oC for 24h followed by centrifugation. Bottom Fraction (BF)74 Fresh H. brasiliensis latex RRIM600 was tapped in ice-chilled flasks. (first 10 mins discarded to minimize contaminants) The latex was filtered through a muslin cloth to remove some coagulants and then centrifuged at 49,000 g for 45min at 4oC. The bottom yellowish fraction is the BF C-serum74 During centrifugation, the clear middle phase that contains aqueous proteins and water present in the cytosol. Washed rubber particle (WRP) Enzymatically active rubber particles from H. brasiliensis (RRIM600) latex were purified by washing four times in 100 mM Tris-HCI (pH 7.5), 5 mM DTT and 0.1 mM AEBSF, centrifugation and suspension in wash buffer. After four washes, 10% glycerol was added and then the WRPs were transferred into liquid nitrogen.

205

Table B.4.4.2 - 1H NMR spectral data of nerol. Position δH (ppm) J (Hz) a 4.07 (d) 12 b 5.43 (t) 27 c’ 1.72 (s) d 2.02 (t) e 2.13 (t) f 5.06 (t) 27 g’ 1.59 (s) h 1.67 (s)

Table B.4.4.3 - 1H NMR spectral data of PN (product 2). Position δH (ppm) J (Hz) a 4.12 (d) 12 b 5.41 (t) 27 c’ 1.74 (s) d 2.02 (t) e 2.13 (t) f 5.09 (t) 27 g’ 1.59 (s) h 1.67 (s) i 4.61 (t) 9 j 3.50 (d) k 1.40~1.80 (m) l 1.40~1.80 (m) m 1.40~1.80 (m)

206

Table B.4.4.4 - 13C NMR spectral data of PN (product 2). Position δC (ppm) A 62 B 124 C 141 C’ 26 D 33 E 26 F 121 G 132 G’ 17 H 24 I 98 J 64 K 27 L 19 M 28

Table B.4.4.5 - 1H NMR spectral data of PN-OH (product 3). Position δH (ppm) J (Hz) a 4.12 (d) 18 b 5.41 (t) n/a c’ 1.74 (s) d 2.14 (t) n/a e 2.14 (t) n/a f 5.36 (t) n/a g 1.59 (s) g’ 1.67 (s) h 3.95 (d) 7 i 4.63 (t) 24 j 3.55 (d) 75 k 1.40~1.80 (m) l 1.40~1.80 (m) m 1.40~1.80 (m) *peaks b,f and d,e are overlapped.

207 Table B.4.4.6 - 13C NMR spectral data of PN-OH (product 3). Position δC (ppm) A 62 B 125 C 141 C’ 23 D 32 E 26 F 122 G 136 G’ 15 H 68 I 98 J 64 K 27 L 19 M 30

Table B.4.4.7 - 1H NMR spectral data of PN-Ts (product 4). Position δH (ppm) J (Hz) a 4.12 (d) 18 b 5.41 (t) 9 c’ 1.74 (s) d 2.14 (t) n/a e 2.14 (t) n/a f 5.36 (t) 9 g 1.59 (s) g’ 1.67 (s) h 4.11 (s) i 4.63 (t) 24 j 3.55 (d) 75 k 1.40~1.80 (m) l 1.40~1.80 (m) m 1.40~1.80 (m) n 7.87 (d) 10 o 7.25 (d) 12 p 2.42 (s) *peaks d,e are overlapped.

208 Table B.4.4.8 - 13C NMR spectral data of PN-Ts (product 4). Position δC (ppm) A 62 B 125 C 141 C’ 23 D 32 E 26 F 122 G 136 G’ 15 H 71 I 97 J 64 K 27 L 19 M 30 N 128 N’ 138 O 132 O’ 144 P 36

Table B.4.4.9 - 1H NMR spectral data of starting PIB-OH. Position δH (ppm) J (Hz) 1 7.35 (m) 2 7.35 (m) 3 7.35 (m) 5 1.94 (d) 36 9 1.43 10 1.12 13 1.56 14 1.94 15 4.63 17 4.60 (d) 40

209 Table B.4.4.10 - 13C NMR spectral data of starting PIB-OH (#16). Position δC (ppm) 1 124 2 129 3 126 4 148 5 71 7 54 8 31 8’ 43 9 55 10 32 10’ 22 11 48 12 32 13 59 14 34 14’ 63 16 27 17 113 18 146

210 Table B.4.4.11 - 1H NMR spectral data of PIB-PN (product 5). Position δH (ppm) J (Hz) a 4.12 (d) 18 b 5.41 (t) 9 c’ 1.74 (s) d 2.14 e 2.14 f 5.36 (t) h 3.71 (s) i 5.18 (t) 12 j 3.47 (d) 24 k 1.40~1.80 (m) l 1.40~1.80 (m) m 1.40~1.80 (m) 1 7.35 (m) 2 7.35 (m) 3 7.35 (m) 5 3.43 (s) 9 1.43 10 1.12 13 1.56 14 1.94 15 4.63 17 4.62 (d) 40

211 Table B.4.4.12 - 13C NMR spectral data of PIB-PN (product 5). Position δC (ppm) A 62 B 125 C 141 C’ 23 D 32 E 26 F 122 G 136 G’ 15 H 73 I 97 J 64 K 27 L 19 M 30 1 124 2 129 3 126 4 148 5 75 7 54 8 31 8’ 43 9 55 10 32 10’ 22 11 48 12 32 13 59 14 34 14’ 63 16 27 17 113 18 146

212 Table B.4.4.13 - 1H NMR spectral data of PIB-Nerol (product 6) Position δH (ppm) J (Hz) a 3.91 (d) 15 b 5.41 (t) 9 c’ 1.74 (s) d 2.14 e 2.14 f 5.36 (t) h 3.71 (s) i 5.18 (t) 12 j 3.47 (d) 24 k 1.40~1.80 (m) l 1.40~1.80 (m) m 1.40~1.80 (m) 1 7.35 (m) 2 7.35 (m) 3 7.35 (m) 5 3.43 (s) 9 1.43 10 1.12 13 1.56 14 1.94 15 4.63 17 4.62 (d) 40

213 Table B.4.4.14 - 13C NMR spectral data of PIB-Nerol (product 6). Position δC (ppm) A 65 B 125 C 141 C’ 23 D 32 E 26 F 122 G 136 G’ 15 H 73 1 124 2 129 3 126 4 148 5 75 7 54 8 31 8’ 43 9 55 10 32 10’ 22 11 48 12 32 13 59 14 34 14’ 63 16 27 17 113 18 146

214 Table B.4.4.15 - 1H NMR spectral data of PIB-Nerol-Br (product 7). Position δH (ppm) J (Hz) a 3.91 (s) b 5.41 (t) 9 c’ 3.9 (s) d 2.14 e 2.14 f 5.36 (t) h 3.71 (s) i 5.18 (t) 12 j 3.47 (d) 24 k 1.40~1.80 (m) l 1.40~1.80 (m) m 1.40~1.80 (m) 1 7.35 (m) 2 7.35 (m) 3 7.35 (m) 5 3.43 (s) 9 1.43 10 1.12 13 1.56 14 1.94 15 4.63 17 4.62 (d) 40

215 Table B.4.4.16 - 13C NMR spectral data of PIB-Nerol-Br (product 7). Position δC (ppm) A 28 B 125 C 141 C’ 23 D 32 E 26 F 122 G 136 G’ 15 H 73 1 124 2 129 3 126 4 148 5 75 7 54 8 31 8’ 43 9 55 10 32 10’ 22 11 48 12 32 13 59 14 34 14’ 63 16 27 17 113 18 146

216 Table B.4.4.19 - 1H NMR spectral data of PN-DVA (product 9). Position δH (ppm) J (Hz) a 3.91 (d) b 5.41 (t) 9 c’ 3.9 (s) d 2.14 e 2.14 f 5.36 (t) h 3.71 (s) g’ 4.46 i 4.58 (t) 12 j 3.47 (d) 24 k 1.40~1.80 (m) l 1.40~1.80 (m) m 1.40~1.80 (m) p 2.31(t) q 2.37 r 1.56 s 2.37 (t) t 7.32 (t) u 4.90 (d) 18

Table B.4.4.46 - 1H NMR spectral data of Nerol-PP-DVA-TMP (product 16). Position δH (ppm) J (Hz) a 4.79 (d) b 5.42 (t) 9 c’ 3.9 (s) d 2.13 e 2.13 f 5.42 (t) h 3.71 (s) g’ 1.76 o 2.37 (t) 12 p 1.60~1.80 (m) q 1.60~1.80 (m) r 2.33 (t) 1 4.08 (t) 2 1.65 3 1.76 4’ 1.12 (s) 5 1.42 (s) 7 0.97 (s)

217 Table B.4.4.47 - 13C NMR spectral data of Nerol-PP-DVA-TMP (product 16). Position δC (ppm) A 62 B 125 C 134 C’ 23 D 32 E 26 F 122 G 136 G’ 15 H 71 1 68 2 37 3 22 4 21 5 71 7 28 N 172 O 42 P 21 Q 13 R 40 S 170 W 60 X 28 Y 22 Z 13

218