Total Synthesis of Isotope-Labeled Isolevuglandins

TOTAL SYNTHESIS OF ISOTOPE-LABELED ISOLEVUGLANDINS

YUNFENG XU

Submitted in partial fulfillment of the requirements for the degree of

Doctor of Philosophy

Thesis Advisor:

Dr. Robert G. Salomon

Department of Chemistry

CASE WESTERN RESERVE UNIVERSITY

Augest 2012

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the dissertation of

Yunfeng Xu

candidate for the Doctor of Philosophy degree *.

(signed) Dr. Michael Zagorski (chair of the committee)

Dr. Anthony Pearson

Dr. Gregory Tochtrop

Dr. Irina Pikuleva

Dr. Robert Salomon

(date) 06/28/2012

* We also certify that written approval has been obtained for any

propriety material contained therein.

This thesis is dedicated to my parents and my aunt.

iii

TABLE OF CONTENTS

Table of Contents ...... iv List of Tables ...... vi List of Figures ...... vi List of Schemes ...... xii Acknowledgements ...... xiv List of Abbreviations and Acronyms ...... xvi Abstract...... xix

Total Synthesis of Isotope-labeled Isolevuglandins

Chapter 1 Introduction ...... 1 1.1 Enzymatic pathways ...... 2 1.2 The free radical pathway and nomenclature of isolevuglandins ...... 5 1.3 Isolevuglandins covalently modify biomolecules ...... 7 1.4 A new strategy of preparation of quantification internal standards ...... 10 1.5 The synthesis of isoLGs ...... 11 1.6 References ...... 14

Chapter 2 Total Synthesis of Deuterium-labeled iso[4]LGE2-d6 ...... 21 2.1 Background ...... 22 2.2 Results and discussion ...... 23

2.2.1 An Improved synthesis of iso[4]LGE2 ...... 23

2.2.2 Synthetic design of deuterium-labeled iso[4]LGE2-d6...... 26

2.2.3 Ring opening of THF-d8 ...... 28 2.2.4 Alkylation of diethyl phosphonoacetone ...... 29

2.2.5 Formation of the top chain: TBDMS deprotection, oxidation and esterification ...... 30

2.2.6 Construction of iso[4]LGE2-d6 carbon skeleton ...... 32

2.2.7 Generationof iso[4]LGE2-d6 acetal: deprotection of the TBDMS silyl ether and hydrolysis of the methyl ester ...... 34

2.2.8 Generation of iso[4]LGE2-d6 ...... 35 2.3 Conclusions ...... 37 2.4 Experimental ...... 38 2.5 References ...... 51

Chapter 3 Total Synthesis of Deuterium-labeled isoLGE2-d6 ...... 55 3.1 Background ...... 56 3.2 Results and discussion ...... 57 3.2.1 Alkylation of propargyl alcohol ...... 59 3.2.2 Acetylation of alkylated propargyl alcohol (3.1) ...... 61 3.2.3 Generation of 3.3: deprotection, oxidation and esterification of 3.2 ...... 62 3.2.4 Selective hydrogenation of the triple bond to a cis double bond using P-2 nickel boride ...... 63 3.2.5 The Appel reaction: bromination of 3.4 to generate the top side chain precursor 3.5 ...... 64

3.2.6 Generation of enone-d6 (3.7) ...... 64

3.2.7 Elaboration of isoLGE2-d6 (3.12) from enone-d6 3.7 ...... 65 3.3 Conclusions ...... 67 3.4 Experimental ...... 68 3.5 References ...... 83

Appendix ...... 86 Bibliography ...... 142

LIST OF TABLES

Table 2.1 The yield of alkylation of propargyl alcohol with 2.1 using different 61

bases

LIST OF FIGURES

Fig. 1.1 The origin of "levuglandin" 4

Fig. 1.2 Strategy for isotope-labeling in previous studies of iso[4]LGE2 10

modification

Fig. 1.3 Phospholipase cleavage sites. An enzyme that displays both PLA1 11

and PLA2 activities is called a Phospholipase B (PLB).

1 2 13 Fig 2.1 H, H and C NMR spectra of TBDMS protected 4-iodobutyl-d8 29

alcohol

Fig 2.2 Determination of the ratio of cis- and trans- isomers of enone-d6 2.5 33

by 1H-NMR

Fig 2.3 Estimation of the yield of hydrolysis of acetal 2.10 to generate 36

iso[4]LGE2-d6

1 Fig. A1 H-NMR spectrum of tert-butyl(1,1,2,2,3,3,4,4-d8-4- 87

iodobutoxy)dimethylsilane (2.1)

2 Fig. A2 H-NMR spectrum of tert-butyl(1,1,2,2,3,3,4,4-d8-4- 88

iodobutoxy)dimethylsilane (2.1)

13 Fig. A3 C-NMR spectrum of tert-butyl(1,1,2,2,3,3,4,4-d8-4- 89

iodobutoxy)dimethylsilane (2.1)

1 Fig. A4 H-NMR spectrum of diethyl(4,4,5,5,6,6,7,7-d8-7-((tert- 90

butyldimethylsilyl)oxy)-2-oxoheptan-3-yl)phosphonate (2.2)

2 Fig. A5 H-NMR spectrum of diethyl(4,4,5,5,6,6,7,7-d8-7-((tert- 91

butyldimethylsilyl)oxy)-2-oxoheptan-3-yl)phosphonate (2.2)

13 Fig. A6 C-NMR spectrum of diethyl(4,4,5,5,6,6,7,7-d8-7-((tert- 92

butyldimethylsilyl)oxy)-2-oxoheptan-3-yl)phosphonate (2.2)

1 Fig. A7 H-NMR spectrum ofdiethyl(4,4,5,5,6,6,7,7-d8-7-hydroxy-2- 93

oxoheptan-3-yl)phosphonate (2.3)

2 Fig. A8 H-NMR spectrum ofdiethyl(4,4,5,5,6,6,7,7-d8-7-hydroxy-2- 94

oxoheptan-3-yl)phosphonate (2.3)

13 Fig. A9 C-NMR spectrum ofdiethyl(4,4,5,5,6,6,7,7-d8-7-hydroxy-2- 95

oxoheptan-3-yl)phosphonate (2.3)

1 Fig. A10 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-(diethoxyphosphono)- 96

6-oxoheptanoate (2.4)

2 Fig. A11 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-(diethoxyphosphono)- 97

6-oxoheptanoate (2.4)

13 Fig. A12 C-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-(diethoxyphosphono)- 98

6-oxoheptanoate (2.4)

1 Fig. A13 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-acetyl-7,7- 99

dimethoxyhept-5-enoate(2.5)

2 Fig. A14 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-acetyl-7,7- 100

dimethoxyhept-5-enoate(2.5)

vii

13 Fig. A15 C-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-acetyl-7,7- 101

dimethoxyhept-5-enoate(2.5)

1 Fig. A16 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-9- 102

(tert-butyldimethylsilyloxy)-6-(dimethoxymethyl)heptadeca-7,11-

dienoate (2.8)

2 Fig. A17 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-9- 103

(tert-butyldimethylsilyloxy)-6-(dimethoxymethyl)heptadeca-7,11-

dienoate (2.8)

1 Fig. A18 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6- 104

(dimethoxymethyl)-9-hydroxyheptadeca-7,11-dienoate (2.9u)

2 Fig. A19 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6- 105

(dimethoxymethyl)-9-hydroxyheptadeca-7,11-dienoate (2.9u)

1 Fig. A20 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6- 106

(dimethoxymethyl)-9-hydroxyheptadeca-7,11-dienoate (2.9l)

2 Fig. A21 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6- 107

(dimethoxymethyl)-9-hydroxyheptadeca-7,11-dienoate (2.9l)

1 Fig. A22 H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6- 108

(dimethoxymethyl)-9-hydroxyheptadeca-7,11-dienoic acid (2.10)

2 Fig. A23 H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6- 109

(dimethoxymethyl)-9-hydroxyheptadeca-7,11-dienoic acid (2.10)

1 Fig. A24 H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6-formyl-9- 110

hydroxyheptadeca-7,11-dienoic acid (2.11)

2 Fig. A25 H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6-formyl-9- 111

viii

hydroxyheptadeca-7,11-dienoic acid (2.11)

1 Fig. A26 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert- 112

butyldimethylsilyl)oxy)hept-2-yn-1-ol (3.1)

2 Fig. A27 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert- 113

butyldimethylsilyl)oxy)hept-2-yn-1-ol (3.1)

13 Fig. A28 C-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert- 114

butyldimethylsilyl)oxy)hept-2-yn-1-ol (3.1)

1 Fig. A29 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert- 115

butyldimethylsilyl)oxy)hept-2-yn-1-yl acetate (3.2)

2 Fig. A30 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert- 116

butyldimethylsilyl)oxy)hept-2-yn-1-yl acetate (3.2)

13 Fig. A31 C-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert- 117

butyldimethylsilyl)oxy)hept-2-yn-1-yl acetate (3.2)

1 Fig. A32 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-ynoate 118

(3.3)

2 Fig. A33 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-ynoate 119

(3.3)

13 Fig. A34 C-NMR spectrum of methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-ynoate 120

(3.3)

1 Fig. A35 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5- 121

enoate (3.4)

2 Fig. A36 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5- 122

enoate (3.4)

13 Fig. A37 C-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5- 123

enoate (3.4)

1 Fig. A38 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-bromohept-5- 124

enoate (3.5)

2 Fig. A39 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-bromohept-5- 125

enoate (3.5)

1 Fig. A40 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-8- 126

(diethoxyphosphoryl)-9-oxodec-5-enoate (3.6)

2 Fig. A41 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-8- 127

(diethoxyphosphoryl)-9-oxodec-5-enoate (3.6)

1 Fig. A42 H-NMR spectrum of (5Z)-methyl 2,2,3,3,4,4-d6-8-acetyl-10,10- 128

dimethoxydeca-5,8-dienoate (3.7)

2 Fig. A43 H-NMR spectrum of (5Z)-methyl 2,2,3,3,4,4-d6-8-acetyl-10,10- 129

dimethoxydeca-5,8-dienoate (3.7)

1 Fig. A44 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12- 130

((tert-butyldimethylsilyl)oxy)-9-(dimethoxymethyl)heptadeca-5,10-

dienoate (3.9u)

2 Fig. A45 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12- 131

((tert-butyldimethylsilyl)oxy)-9-(dimethoxymethyl)heptadeca-5,10-

dienoate (3.9u)

1 Fig. A46 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12- 132

((tert-butyldimethylsilyl)oxy)-9-(dimethoxymethyl)heptadeca-5,10-

dienoate (3.9l)

2 Fig. A47 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12- 133

((tert-butyldimethylsilyl)oxy)-9-(dimethoxymethyl)heptadeca-5,10-

dienoate (3.9l)

1 Fig. A48 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9- 134

(dimethoxymethyl)-12-hydroxyheptadeca-5,10-dienoate (3.10u)

2 Fig. A49 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9- 135

(dimethoxymethyl)-12-hydroxyheptadeca-5,10-dienoate (3.10u)

1 Fig. A50 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9- 136

(dimethoxymethyl)-12-hydroxyheptadeca-5,10-dienoate (3.10l)

1 Fig. A51 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9- 137

(dimethoxymethyl)-12-hydroxyheptadeca-5,10-dienoate (3.10l)

1 Fig. A52 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9- 138

(dimethoxymethyl)-12-hydroxyheptadeca-5,10-dienoic acid

(isoLGE2-d6 acetal, 3.11)

1 Fig. A53 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9- 139

(dimethoxymethyl)-12-hydroxyheptadeca-5,10-dienoic acid

(isoLGE2-d6 acetal, 3.11)

1 Fig. A54 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9-formyl-12- 140

hydroxyheptadeca-5,10-dienoic acid (isoLGE2-d6, 3.12)

2 Fig. A55 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9-formyl-12- 141

hydroxyheptadeca-5,10-dienoic acid (isoLGE2-d6, 3.12)

LIST OF SCHEMES

Scheme 1.1 Enzymatic pathways of the oxidation of AA-PC 3

Scheme 1.2 Rearrangement of the endoperoxide to generate γ-ketoaldehyde 5

Scheme 1.3 Free radical induced autoxidation of AA-PC to generate isoLGs 6

Scheme 1.4 Modification of primary amine group by isoLGs to generate 8

pyrrole and lactam/hydroxylactam

Scheme 1.5 Synthesis of internal standard iso[4]LGE2-d31-lysoPE- 9

lactam/HL

Scheme 1.6 PLD hydrolysis of isoLG-modified PEs 11

Scheme 1.7 Synthesis of isoLG-d6-ethanolamine-lactam/HL 11

Scheme 2.1 Original synthesis of iso[4]LGE2 23

Scheme 2.2 Improved synthesis of iso[4]LGE2 24

Scheme 2.3 A previous synthesis of tributylstannyl side chain intermediate 25

2.6

Scheme 2.4 Improved synthesis of tributylstannyl side chain 26

Scheme 2.5 Synthesis of iso[4]LGE2-d6 27

Scheme 2.6 Ring opening of THF-d8 28

Scheme 2.7 Alkylation of diethyl phosphonoacetone 29

Scheme 2.8 TBDMS deprotection of 2.2 30

Scheme 2.9 Formation of the top chain intermediate 2.4 for iso[4]LGE2-d6 31

Scheme 2.10 Conversion of phosphonate-d6 2.4 to enone-d6 2.5 32

Scheme 2.11 Formation of the carbon skeleton of iso[4]LGE2-d6 33

xii

Scheme 2.12 TBDMS deprotection of 2.8 34

Scheme 2.13 Generation of iso[4]LGE2-d6 acetal 35

Scheme 2.14 Hydrolysis of dimethyl acetal 2.10 to generate iso[4]LGE2-d6 35

2.11

Scheme 3.1 Synthetic strategies of bromoalkenoate in previous synthesis 58

Scheme 3.2 Synthesis top side chain precursor (3.5) of isoLGE2-d6 59

Scheme 3.3 Alkylation of propargyl alcohol 60

Scheme 3.4 Acetylation of alkylated propargyl alcohol (3.1) to give 3.2 62

Scheme 3.5 Generation of 3.3: oxidation, deprotection and esterification of 63

3.2

Scheme 3.6 P-2 nickel hydrogenation of 3.3 63

Scheme 3.7 The Appel reaction 64

Scheme 3.8 Generation of enone-d6 3.7 64

Scheme 3.9 From enone-d6 3.7 to isoLGE2-d6 3.12 65

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ACKNOWLEDGEMENTS

First and foremost, I want to express my genuine gratitude and respect to my research advisor, Dr. Robert G. Salomon. He provided me the best possible facilities, opportunities and challenging tasks which build a solid foundation for my future goals.

Without his invaluable advice, I would not have been able to present this thesis. My research aptitudes have been tremendously enhanced over the years under his guidance. I also feel very grateful for his recommendation to help me to get my job.

I would also like to thank my committee members, Dr. Michael Zagorski, Dr.

Anthony Pearson, and Dr. Gregory Tochtrop for their valuable time and input on my thesis. Special thanks to Dr. Irina Pikuleva for agreeing to be on my committee and for her precious time and input on my thesis.

I would like to thank Dr. James M. Laird. Jim was there the first day when I joined Salomon Group. I still remember he helped me cleaning my office desk when I first came to the lab. He was there throughout my studying and always willing to help whenever I had questions or problems. He taught me a lot about organic synthesis. I really enjoyed discussion with him about research as well as movies.

I would like to thank my labmates, Dr. Mikhail Linetsky, Hua Wang, Yu Zhang,

Yalun Cui, Wenyuan Yu, Wenzhao Bi, Junhong Guo, Guangyin Wang, Dawit

Ghebrehiwot, Nicholas Tomko and my former colleagues Dr. Liang Lu, Dr. Wujuan

Zhang, Dr. Xi Chen, Dr. Wei Li, Dr. Xiaodong Gu, Dr. Jaewoo Choi, Dr. Li Hong, Dr.

xiv

Xiaoxia Zhang, Dr. Detao Gao, Rong Zhou, Liang Xin, Yuanyuan Qian, for their

friendship, which has made my life and study very enjoyable.

I would like to thank Dr. Dale Ray and Dr. Jim Faulk for the training and help on

NMR and Mass Spectrometry. I would like to thank all the current staff members and previous staff members Zedeara Diaz and Pat Eland in the Department of Chemistry for their help on numerous occasions.

Last but absolutely not the least, I would like to thank my parents and my aunt for their endless love, unconditional support and countless sacrifices throughout the course of my education.

LIST OF ABBREVIATIONS AND ACRONYMS

AA arachidonic acid

AA-PC 2-arachidonyl-phosphatidylcholine

AcOH acetic Acid

AMD age related macular degeneration b broad n-BuLi n-butyllithium

Bu3Sn tributylstannyl

CDCl3 deuterated chloroform

13C-NMR Carbon magnetic resonance

COX cyclooxygenase d62-dP-PE 1,2-dipalmitoyl- d62-sn-glycero-3-phosphoethanolamine

DMP Dess-Martin periodinane

DNA deoxyribonucleic acid

EA ethyl acetate

Et ethyl

Et2O diethyl ether

EtOH ethanol

H hexanes

HL hydroxylactam

4-HNE 4-hydroxynonenal

1H-NMR Proton magnetic resonance

2H-NMR Deuterium magnetic resonance

xvi

15-HPETE 15-hydroperoxy-eicosatetraenoate

HRMS high resolution mass spectrometry

Hz hertz iso[4]LGE2 iso[4]levuglandin E2 isoK isoketal isoLG isolevuglandin isoLGD2 isolevuglandin D2 isoLGE2 isolevuglandin E2 isoP isoprostane isoPGH2 isoprostane endoperoxide

J coupling constant

LDL low-density lipoprotein

LG levuglandin

LGD2 levuglandin D2

LGE2 levuglandin E2

Li2Me2CuCN lithium dimethylcyanocuprate lyso-PE 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine

MC methylene chloride

Me methyl

MeOH methanol

MHz megahertz

MS mass spectrometry

NMR nuclear magnetic resonance

PC phosphatidylcholine

xvii

PG prostaglandin

PGD2 prostaglandin D2

PGE2 prostaglandin E2

PGG2 prostaglandin endoperoxide G2

PGH2 prostaglandin endoperoxide H2

PLA2 phospholipase A2

PLD phospholipase D ppm parts per million

PUFA polyunsaturated fatty acid

Rf retention factor

RT room temperature s singlet t triplet

TBAF tert-butylammonium fluoride

TBDMS tert-butyldimethylsilyl

TBDMSCl tert-butyldimethylsilyl chloride tBu tert-butyl tBuLi tert-butyllithium

THF tetrahydrofuran

TLC thin layer chromatography

TMS trimethylsilyl

xviii

Total Synthesis of Isotope-labeled Isolevuglandins

Abstract

YUNFENG XU

Oxidation of arachidonyl-phosphotidylcholine (AA-PC) follows two different

major pathways: an enzymatic pathway and a free radical pathway (the isoprostane

pathway). The enzymatic pathway generates enantiomerically pure levuglandin E2 (LGE2) by a rearrangement of the prostaglandin endoperoxide PGH2. Free radical induced

autoxidation AA-PC generates isolevuglandins (isoLGs) as a mixture of racemic

structural and stereo isomers of LGE2. The γ-ketoaldehyde functionality of LGs and

isoLGs makes them extraordinarily reactive toward primary amino groups in

biomolecules to form covalent LG/isoLG adducts. Because they accumulate, this makes

them attractive as dosimeters to evaluate oxidative injury in tissues. In the previous studies of isoLG modifications to protein and ethanolamine phospholipid, the isotope- labeled internal standards have been prepared from the targets to which iso[4]LGE2

covalently binds. This method of preparing internal standards greatly limits the progress

of research since for every modification an isotope-labeled target molecule must be

prepared. However if isotope-labeled isoLGs were available, they could be used

universally for the preparation of internal standards for any isoLG-modified adduct.

Unlabeled iso[4]LGE2 and isoLGE2 were synthesized previously. In the syntheses

of deuterated iso[4]LGE2-d6 and isoLGE2-d6, the six deuterium atoms were introduced in

xix the top side chains. We could not follow the same strategy as used for syntheses of unlabeled iso[4]LGE2 and isoLGE2 since the deuterated substitutes of the starting materials used for the synthesis of top side chains in the previous syntheses are either not commercially available or very expensive. In our syntheses, we developed variations of previous syntheses that allowed the use of an inexpensive and readily starting material,

THF-d8, to prepare deuterated iso[4]LGE2-d6 and isoLGE2-d6. The method for the synthesis of deuterated isoLGE2-d6 is also applicable for the synthesis of unlabeled isoLGE2 from THF, and is more efficient than the previous method.

The syntheses did not generate pure diastereomers as in the previous syntheses.

This is satisfactory since all diastereomers are produced through the free radical oxidation of arachidonic acid and it was not desirable to only generate one diastereomer.

Chapter 1

Introduction

Lipids are essential cell membrane components, and incorporate abundant

polyunsaturated fatty acids (PUFAs) that are particularly susceptible to oxidative damage.

Lipids play important roles in many biological functions, such as in energy storage, in signaling pathways,1, 2 and in regulation of gene expression.3, 4

Lipid oxidation, although important to human health in biosynthesis of

prostaglandins, thromboxanes, prostacyclins and leukotrienes, etc., is believed to

contribute to many diseases, such as amyotrophic lateral sclerosis,5 age-related macular

degeneration (AMD),6, 7 end-stage renal disease (RD),8, 9 Alexander’s,10 Alzheimer’s,11-13 and Parkinson’s14 diseases. It is also suspected to participate in the pathogenesis of

atherosclerosis (AS),9, 15, 16 antiphospholipid antibody syndrome,17 rheumatoid arthritis,18 inflammatory bowel disease19 and multiple sclerosis.20, 21

Arachidonic acid (AA), a linear twenty carbon methylene interrupted polyunsaturated fatty acid, is linked as an ester to 2-lysophosphotidylcholine in arachidonyl-phosphotidylcholine (AA-PC), one of the most abundant phospholipids in human low density lipoprotein (LDL). It may be oxidized through two different major pathways: an enzymatic pathway and a free radical pathway.

1.1 Enzymatic pathways

The enzyme phospholipase A2 (PLA2) converts the membrane-bound AA-PC to

free AA. Oxidation of AA can be catalyzed by cyclooxygenase (COX) and lipoxygenase

(LOX) (Scheme 1.1). Lipoxygenase converts AA into a fatty acid hydroperoxide by

inserting one molecule of diatomic oxygen to form 15-hydroperoxy-eicosatetraenoate

(15-HPETE).

(CH2)3COO-PC C5H11

AA-PC

PLA2

(CH2)3COOH O (CH2)3COOH (CH2)3COOH C5H11 C H O C5H11 COX LOX 5 11 AA OOH OOH 15-HPETE PGG2

hydroperoxidase

(CH2)3COOH O

O C5H11

OH PGH2

O HO (CH ) COOH (CH2)3COOH 2 3 (CH2)3COOH

C H C5H11 5 11 C5H11 O HO OH OH O OH TXA2 PGE2 PGD2

(CH2)3COOH

O O

(CH2)3COOH (CH2)3COOH

C5H11 C5H11 C5H11

O OH O OH OH OH LGE LGD2 2 PGI2

Scheme 1.1 Enzymatic pathways of the oxidation of AA-PC

In the cyclooxygenase pathway, a phospholipase actively cleaves AA from

membrane phospholipids. After the free AA moves through a channel to the second

catalytic site, two molecules of diatomic oxygen are chemoselectively and

stereospecifically added to one molecule of AA to produce a highly reactive

prostaglandin endoperoxide G2 (PGG2), which is quickly reduced by hydroperoxidase to

o produce the unstable prostaglandin endoperoxide H2 (PGH2, t1/2 = 5 min at 37 C in

22 aqueous solution). PGH2 can undergo enzymatic and non-enzymatic (solvent induced)

rearrangements to generate an array of physiologically active molecules such as

prostaglandin E2 (PGE2), or prostaglandin D2 (PGD2), prostaglandin I2 (PGI2),

thromboxane A2 (TXA2).

About 30 years ago Salomon et al. first reported a novel alternative rearrangement

of PGH2 to produce two γ-ketoaldehydes which were named levuglandin E2 (LGE2) and

23, 24 levuglandin D2 (LGD2). The word levuglandin comes from the combination of

levulinaldehyde and prostaglandin since levuglandins are derivatives of levulinaldehyde with prostaglandin side chains (Fig. 1.1).25

Fig. 1.1 The origin of "levuglandin"

Detailed studies revealed that the rearrangement of the endoperoxide to generate

the γ-ketoaldehyde was a result of a 1,2 hydride shift during the cleavage of three bonds

in a concerted mechanism (Scheme 1.2) with a polarized transition state in aqueous solution.26, 27

1.2 The free radical pathway and nomenclature of isolevuglandins

Unlike the enzymatic pathways which convert free AA to an endoperoxide

intermediate, the free radical pathway oxidizes AA-PC directly, in preference to free

AA,25 to produce stereo and structural isomers of prostaglandins28 which are now called

isoprostanes.29 This is why the free radical pathway is also called the isoprostane pathway.

Isoprostanes are formed from rearrangement of isoPGH2-PC. As levuglandins can be

produced via rearrangement of PGH2, isolevuglandins (isoLGs), stereo and structural

25 isomers of levuglandins, are formed via rearrangement of isoPGH2 (Scheme 1.3). And

because abstraction of a hydrogen atom occurs nonregioselectively at any doubly allylic

methylene, the free radical pathway not only produces a stereoisomeric mixture of

levulinaldehyde derivatives with PG side chains, i.e., isoLGs, but also a new group of levulinaldehyde derivatives with non-prostanoid side chains, which, proposed by Dr.

Salomon,25 are collectively called iso[n]LGs, where the bracketed interger indicates the

number of carbon atoms of the carboxylic side chain. If in an isoLG molecule the acetyl

5 substituent is nearest to the carboxylic side chain, it is designated as E series; if the formyl group is nearest to the carboxylic side chain, it is designated as D series.

The terms “isoketals” or “isoKs” were alternatively used in the original isoLG

nomenclature to “distinguish them from levuglandins formed by rearrangement of the

30 cyclooxygenase endoperoxide intermediates, PGH2”. However, this nomenclature is

erroneous because isoLGs are not ketals. Furthermore, the exact same levuglandin

molecules, LGE2 and LGD2, are generated by both the cyclooxygenase and isoprostane

pathways. The difference between these two pathways is that a racemic mixture of stereo

and structural isomers is cogenerated with LGE2 and LGD2, whereas the cyclooxygenase

pathway generates only a single enantiomer of each product. On the other hand, the

isoLG nomenclature can easily distinguish the difference between stereoisomers of LGs

and structural isomers of LGs. For example, LGE2, one of the stereoisomers designated collectively as isoLGE2, is a product of both the COX and isoprostane pathways, whereas

all other stereoisomers of isoLGE2 are only produced through the isoprostane pathway.

1.3 Isolevuglandins covalently modify biomolecules

Because of their γ-ketoaldehyde functionality, LGs and isoLGs are extraordinarily

reactive toward primary amino groups to covalently modify protein, DNA31 and

phosphatidylethanolamine (PE)32 to generate an aromatic pyrrole derivative. This transformation, a Paal-Knorr synthesis of pyrroles, proceeds through a Schiff base and its enamine tautomer. Since LG-derived pyrroles are electron-rich, they are readily oxidized by air to form relatively stable end products, lactams and hydroxylactams (Scheme

1.4).33, 34 It was found35-38 that these modifications are closely related to age-related

macular degeneration (AMD), atherosclerosis (AS) and cardiovascular disease (CVD).

The mitochondrial enzyme cytochrome P450 27A1 (CYP27A1) plays an important role in the maintenance of cholesterol homeostasis, bile acid biosynthesis, and

39 activation of vitamin D3 by catalyzing the C27-hydroxylation of cholesterol and other sterols. Recently CYP27A1 was found to be expressed in the retina40, 41 and shown to be the major contributor to enzymatic degradation of cholesterol.42 A recent study43 found that iso[4]LGE2 treatment of purified recombinant CYP27A1 in solution diminished

15 enzyme activity and resulted in the formation of iso[4]LGE2 adducts. Using a N-labeled

CYP27A1 modified with iso[4]LGE2 as internal standard, the subsequent analysis of proteins extracted from a human retina demonstrated that iso[4]LGE2-modified

CYP27A1 is present in the retina. Future studies will explore the proposition that levels of CYP27A1-isoLG serve as a marker for both oxidative stress and impaired cholesterol elimination.

The primary amino groups of phosphatidylethanolamines (PEs) are also susceptible to covalent modification by lipid peroxidation products. Modifications of PEs

by aldehydes generated in vivo, such as 4-HNE44 and acrolein,45 have been reported.

32 Salomon et al. confirmed that iso[4]LGE2 covalently binds ethanolamine phospholipids

in vitro and in PEs extracted from human plasma and mouse liver to form covalent

pyrrole adducts that are oxidized in air to deliver stable lactam and hydroxylactam (HL)

adducts. For this study, an internal standard was prepared from modification of 1,2-

dipalmitoyl-d62-sn-glycero-3-phosphatidylethanolamine (d62-dP-PE) with iso[4]LGE2.

The resulting pyrrole adduct was converted into lactam/HL under O2 atmosphere,

followed by selective hydrolysis by phospholipase A2 (PLA2) to finally deliver

iso[4]LGE2-d31-lysoPE-lactam/HL as the internal standard for quantification (Scheme

1.5).

CD3(CD2)14COO COOH O O + P NH3 C H O O 5 11 CD3(CD2)14COO O OH dP-PE iso[4]LGE2 X CD3(CD2)14COO O O O2 COOH P N O O C5H11 CD3(CD2)14COO O OH iso[4]LGE2-d62-dP-PE-lactam, X=H iso[4]LGE2-d62-dP-PE-HL, X=OH

X HO O O PLA2 COOH P N O O C5H11 CD3(CD2)14COO O OH iso[4]LGE2-d31-lysoPE-lactam, X=H iso[4]LGE2-d31-lysoPE-HL, X=OH

Scheme 1.5 Synthesis of internal standard iso[4]LGE2-d31-lysoPE-lactam/HL

1.4 A new strategy of preparation of quantification internal standards

For both studies, the isotope-labeled internal standards have been prepared from

the deuterated analogues of target molecules to which iso[4]LGE2 covalently binds (Fig.

1.2). Therefore, for every study requiring quantification of an isoLG modification, an

isotope-labeled target molecule must be prepared. This greatly complicates the research.

However if isotope-labeled isoLGs were available, they could be used universally for the

preparation of internal standards for any isoLG-modified adduct.

Target molecules

15 N-CYP27A1 d62-dP-PE

15 iso[4]LGE2- N-CYP27A1 iso[4]LGE2-d31-lysoPE-lactam/HL

Internal standards

Fig. 1.2 Strategy for isotope-labeling in previous studies of iso[4]LGE2 modification

32 In the above-mentioned iso[4]LGE2-PE study, since PEs with a variety of fatty acid hydrocarbon chains present in vivo will be modified by iso[4]LGE2, detecting the levels of individual species would be difficult and of little value. A key simplifying strategy for this study was to treat the iso[4]LGE2-PE adducts with PLA2 to generate mainly 1-palmitoyl-2-lysoPE-HL from heterogeneous mixtures of phospholipids with a variety of acyl groups on the 2 position. This greatly increased the detection limits of iso[4]LGE2-PE adducts. For the previous preparation of an internal standard, the isotope

was introduced into one of the fatty acid hydrocarbon chains starting from expensive 1,2-

dipalmitoyl-d62-sn-glycero-3-phosphatidylethanolamine (d62-dP-PE) that is no longer

commercially available.

With isotope-labeled isoLGs, a new strategy can be applied for the preparation of

internal standards. As discussed above, in order to enrich iso[4]LGE2-PE adducts, PLA2

was used to selectively hydrolyze all PE adducts into iso[4]LGE2-lysoPE adducts. There

are four major classes of phospholipases, termed A, B, C and D, distinguished by the type

of reaction which they catalyze, as shown in Fig. 1.2. Thus instead of using PLA2, PLD

can be used to hydrolyze the modified phospholipid at the ethanolamine phosphate ester

bond, releasing isoLG-ethanolamine adducts (Scheme 1.6). The corresponding isotope-

labeled internal standard can be prepared accordingly without using any phospholipids

(Scheme 1.7). This strategy should enable the detection of modified PEs with different

acyl chains as a single derivative and would greatly increase the detection limits.

PLA2 PLC PLD

PLA 1

Fig. 1.3 Phospholipase cleavage sites. An enzyme that displays both PLA1 and

PLA2 activities is called a Phospholipase B (PLB).

1.5 The synthesis of isoLGs

Some of the isolevuglandins were successfully synthesized previously.

46 Iso[4]LGE2 was first synthesized by G. Subbanagounder et al. in 1997. This synthesis

suffers from two “bottlenecks”: (1) synthesis of the lower prostanoid side chain and (2)

the use of isopropylene-D-glyceraldehyde as a chiral latent aldehyde necessitating

deprotection of the acetonide and oxidative cleavage of diol to an aldehyde. An improved

synthesis of iso[4]LGE2 developed by J. Laird in Salomon’s group eliminated these

“bottlenecks”. In the improved synthesis, 2,2-dimethoxyacetaldehyde was used to

provide enone intermediates with a dimethyl acetal as a masked aldehyde, which can be

hydrolyzed under mild conditions at the end of the synthesis to generate iso[4]LGE2. This

approach eliminates the additional oxidative step. The use of commercially available cis-

3-nonenol greatly simplified the synthesis of a precursor of lower prostanoid side chain.

The synthesis of a deuterium-labeled iso[4]LGE2-d6 described in this thesis was

accomplished based on Laird’s improved synthesis of iso[4]LGE2. Deuterium was introduced in the top chain using THF-d8 as the source of deuterium.

Previously in the Salomon group, different stereoisomers of LGE2 were

successfully synthesized.47 In this synthesis, the top segment, methyl cis-7-bromohept-5-

enoate, was synthesized via a literature method.48 Another similar synthesis was also

49, 50 reported. Both methods are not applicable for synthesis of deuterated isoLGE2-d6 due to harsh reaction conditions and the expense anticipated for introduction of deuterium.

We developed a more efficient synthetic strategy using readily available starting materials (THF-d8 and propargyl alcohol) for synthesis of the top segment. It is

noteworthy that this method is also applicable for synthesis of non-deuterated isoLGE2 from THF.

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Chapter 2

Total Synthesis of Deuterium-labeled iso[4]Levuglandin E2-d6

2.1 BACKGROUND

As discussed in Chapter 1, iso[4]LGE2 is one of the structural isomers of isolevuglandins generated from free radical induced autoxidation of AA. It was reported

1, 2 that iso[4]LGE2 covalently binds with proteins and ethanolamine phospholipids. For these previous studies, internal standards were prepared from isotope-labeled target molecules (proteins and PEs) that were modified with iso[4]LGE2. This method of preparing internal standards greatly limits the progress of research since for every modification an isotope-labeled target molecule must be prepared. In contrast, deuterium- labeled iso[4]LGE2-d6 can be used universally for preparation of deuterium-labeled internal standards of any iso[4]LGE2 adducts thus greatly facilitates the studies. This chapter presents a method for the synthesis of deuterium-labeled iso[4]LGE2-d6 that is a

3 modification of a previous synthesis of unlabeled iso[4]LGE2. Deuterium was introduced in the top chain using THF-d8 as the source of deuterium.

2.2 RESULTS AND DISCUSSION

2.2.1 An improved synthesis of iso[4]LGE2

Iso[4]LGE2 was first synthesized by G. Subbanagounder et al. in 1997 (Scheme

2.1).4 In this synthesis, isopropylidene D-glyceraldehyde was used as a chiral latent aldehyde to control the stereochemistry α to the aldehyde carbonyl group. This is unnecessary for the isoLGs since they are generated non-stereoselectively as diastereomeric mixtures. Furthermore, two steps are required to deprotect the latent aldehyde to generate iso[4]LGE2: first removal of the acetonide protecting group by heating with dilute acetic acid and then oxidative cleavage of the diol with sodium periodate.

According to a modification of the synthetic strategy for isoLGE2 introduced by

5 Amarnath et al, J. Laird in Salomon’s group improved the synthesis of iso[4]LGE2 by

using 2,2-dimethoxyacetaldehyde to provide enone intermediates with a dimethyl acetal as a masked aldehyde, which can be hydrolyzed under mild conditions at the end of the

synthesis to generate iso[4]LGE2, and this approach eliminates the additional oxidative

step (Scheme 2.2).

A previous synthesis of the tributylstannyl side chain precursor 2.6 requires

stereoselective reduction of an alkyne precursor, which is a tedious and lengthy process

and requires a difficult purification (Scheme 2.3).6 That preparation of the tributylstannyl

side chain started from 1-heptyne, which was converted to a lithium acetylide by n- butyllithium, and the acetylide reacted with 1-bromo-2,2-diethoxyethane to give 1,1- diethoxynon-3-yne. The alkyne was reduced to a cis-alkene and then the diethyl acetal was hydrolyzed by oxalic acid to deliver the aldehyde cis-3-nonenal (2.12). The aldehyde

2.12 was reacted with trans-lithio-trimethylsilyl ethylene to form the racemic alcohol, which could be resolved subsequently. TBDMS protection of the hydroxyl and stannyl desilylation generated the tributylstannyl side chain precursor 2.6.

In an improved synthesis by J. Laird3 (Scheme 2.4), the aldehyde cis-3-nonenal

(2.12) was prepared from commercially available cis-3-nonenol7, 8 by oxidation to cis-3- nonenal (2.12) in one step with excellent yield (90 – 95%) using the Dess-Martin periodinane (DMP).9-11 Because the aldehyde (2.12) is very unstable, it was used

immediately without further purification to react with (E)-lithio-(tri-n-butylstanayl)

ethylene prepared from (E)-1,2-bis(tri-n-butylstannyl)ethylene (2.13) to give the racemic

tributylstannyl alcohol 2.14. This was converted to the tributylstannyl side chain

precursor 2.6 by protecting the alcohol with atert-butyldimethylsilyl group. (E)-1,2-

bis(tri-n-butylstannyl)ethylene (2.13) was produced from tributylstannylacetylene,12 which was generated by the reaction of acetylene with nBuLi in THF followed by tributyltin chloride.13, 14

2.2.2 Synthetic design of deuterium-labeled iso[4]LGE2-d6

A synthesis of deuterium labeled iso[4]LGE2-d6 was designed to exploit features of the previous synthesis of iso[4]LGE2. In the synthesis of iso[4]LGE2-d6, the six deuterium atoms were introduced in the top chain. In the synthesis of iso[4]LGE2, the top

chain is introduced as phosphonate ester that was assembled by the reaction of diethyl(2-

oxopropyl)phosphonate with ethyl 4-iodobutyrate, which is commercially available.

Deuterated ethyl 4-iodobutyrate is not commercially available. Therefore, the synthesis

started with ring opening of THF-d8 to obtain a TBDMS protected 4-iodobutyl-d8 alcohol

(2.1), which was then reacted with diethyl(2-oxopropyl)phosphonate to produce a

TBDMS protected phosphonated alcohol 2.2. The TBDMS protecting group was then

removed by acetic acid and the resulting alcohol was oxidized by Jones reagent to

26 produce a carboxylic acid, which was converted to a methyl ester by reaction with diazomethane to finally deliver the deuterated top chain synthon 2.4 for iso[4]LGE2-d6.

Once the top chain was built up, the subsequent synthetic steps for iso[4]LGE2-d6

(Scheme 2.5) were similar to those of the improved iso[4]LGE2 synthesis (Scheme 2.2).

2.2.3 Ring opening of THF-d8

Ring opening of deuterated THF to generate a TBDMS protected 4-iodobutyl-d8

alcohol 2.1 was achieved by the same procedure as used for ring opening of unlabeled

THF,15 as shown in Scheme 2.6. It would be desirable for the subsequent steps in the

synthesis if THF-d8 were converted to non-protected 4-iodobutyl-d8 alcohol. But the non-

protected 4-iodobutyl-d8 alcohol is very unstable and will easily undergo the reverse

reaction converting the ring opened product back into the starting material. Therefore, it

cannot be stored for long time, and a TBDMS protected alcohol is more favorable. The

formation of 2.1 was confirmed by 1H, 2H and 13C NMR as well as HRMS.

1H-NMR of product 2.1 only showed two signals which come from the three

methyl groups (δ 0.84 ppm) of tert-butyl group and two methyl groups (δ 0.00 ppm) on the silicon of the TBDMS group. 2H-NMR clearly showed four signals which correspond

to the four deuterated methylene groups. Note that the 13C-NMR signals of the deuterated

carbons are very weak, and since 2H was not decoupled, the signal of each deuterated carbon was split by two deuterium atoms resulting in a quintet of peaks in a ratio of

1:2:3:2:1.

1H-NMR 2H-NMR

13C-NMR

1 2 13 Fig 2.1 H, H and C NMR spectra of TBDMS protected 4-iodobutyl-d8 alcohol 2.1

2.2.4 Alkylation of diethyl phosphonoacetone

The alkylation of diethyl(2-oxopropyl)phosphonate with 2.1 was achieved by the same procedure as used for ethyl 4-iodobutyrate in the synthesis of iso[4]LGE2, but in better yield (77% in comparison to 55%),4, 16, 17 as shown in Scheme 2.7. In the 13C-NMR

spectrum of 2.2, only three signals of deuterated carbon were observed. The signal at δ

31.4 ppm was obscured by the signal of the protonated carbon of the carbonyl methyl

group. The yield was calculated based on the starting material 2.1 reacted. Based on the

amount of 2.1 recovered, only 41% of 2.1was converted to product.

2.2.5 Formation of the top chain: TBDMS deprotection, oxidation and esterification

Deprotection of the TBDMS ether to release the free alcohol 2.3 was easily

accomplished by hydrolysis in a solution of acetic/water (3/1, v/v) at room temperature

overnight in excellent yield (94%), as shown in Scheme 2.8. By comparison of the 1H-

NMR spectra of 2.2 and 2.3, the most obvious evidence of formation of 2.3 was the disappearance of signals from the TBDMS group (δ 0.00 ppm, 0.85 ppm) and retention of all the rest of the peaks. Also, in the 13C-NMR spectrum of 2.3, the overlapped peak of

deuterated carbon on 2.2 moved slightly downfield to δ 31.2 ppm.

Conversion of alcohol 2.3 to a carboxylic acid can be achieved by two routes: (1)

two mild oxidations that (i) generate an aldehyde and then (ii) convert it into carboxylic

acid, or (2) a harsher alternative one step oxidation using Jones reagent to directly

convert the alcohol to a carboxylic acid. We used the second route, as shown in Scheme

2.9, since it is more efficient, although it is more challenging to isolate and purify the

product. However, we found that the crude product could be directly used for

esterification without purification.

Methylation of the carboxylic acid was achieved by treatment with freshly

prepared diazomethane. Diazomethane as a methylating agent was first reported in the

late 19th century.18, 19 Nowadays it is widely used for the preparation of simple methyl

esters from the corresponding carboxylic acids. It can conveniently be generated under

mild conditions by treating a suitable precursor20-22 with an alkaline base. Since diazomethane is an extremely toxic, carcinogenic, odorless and explosive yellow gas, not suitable for long time storage, it was prepared freshly when needed. In most of the

literature a distillation/condensation apparatus is used to collect diazomethane solution,

which is unnecessarily complicated. Also because it is an extremely sensitive explosion

when dry, ground glass joints must be avoided. In our lab, diazomethane was generated

by hydrolyzing N-nitrosomethylurea23 in a mixture of 50% aqueous KOH solution

overlayered with diethyl ether in an Erlenmeyer flask with a clean and smooth inner

surface.

The crude product was purified by flash chromatography to give an overall yield

of 70% for the oxidation and methylation. By comparison of the NMR spectra of the

product with those of 2.3, formation of 2.4 can be confirmed with the appearance of a

singlet (δ 3.63 ppm) in the 1H-NMR spectrum of 2.4 corresponding the methyl ester

group, the appearance of a singlet (δ 173.53 ppm) in the 13C-NMR spectrum

corresponding the carbonyl carbon on the ester group as well as the loss of two deuterium

signals in the 2H-NMR spectrum and one deuterated carbon signal in the 13C-NMR spectrum.

2.2.6 Construction of iso[4]LGE2-d6 carbon skeleton

The masked aldehyde 2.5 was synthesized by adding sodium hydride to the phosphonate-d6 2.4 to deprotonate the methane α to the ketone, followed by addition of a

solution of 2,2-dimethoxyglyoxal in ethyl ether to obtain 2.5 (Scheme 2.10) as a mixture

of cis- and trans- isomers in a ratio of 1:4, which is consistent with reported results for

the unlabeled analogue.4 The ratio of cis- and trans- isomers was determined by

comparison of the integrations of either two hydrogens on the carbon-carbon double bond

’ ’ (Ha and Ha ) or two hydrogens on the acetal carbon (Hb and Hb ) of the two isomers (Fig.

2.2).

’ ’ Ha Hb

H H b a

4.01 0.98 3.95 1.10 6.6 6.4 6.2 6.0 5.8 5.6 5.4 5.2 5.0 4.8 f1 (ppm)

9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

1 Fig 2.2 Determination of the ratio of cis- and trans- isomers of enone-d6 2.5 by H-NMR

The carbon skeleton of iso[4]LGE2-d6 was constructed by connecting the

tributylstannyl side chain intermediate 2.6, which was synthesized as reported

previously,3 to enone 2.5 via a cyanocuprate reaction (Scheme 2.11). Cyanocuprates24-30 are important in organic synthesis due to their high and selective reactivity. They react with α,β-unsaturated ketones resulting in conjugate 1,4-addition instead of 1,2-addition to the carbonyl group that occurs when a simple alkyllithium is used.

Addition of two equivalents of methyllithium to copper (I) cyanide resulted in

cyanocuprate Li2Me2Cu(CN), which reacted with the tributylstannyl side chain

intermediate 2.6 via transmetalation to form the higher order vinylcyanocuprate 2.7.

Addition of enones 2.5 to the vinylcyanocuprate 2.7 provided the conjugate addition products 2.8 as a mixture of stereoisomers.

2.2.7 Generation of iso[4]LGE2-d6 acetal: deprotection of the TBDMS silyl ether and

hydrolysis of the methyl ester

The silyl ether of the iso[4]LGE2-d6 carbon skeleton was hydrolyzed with 1.5

equiv of TBAF in THF overnight at room temperature, as shown in Scheme 2.12. TLC

analysis (30% ethyl acetate in hexanes) of the crude product showed that there were two

spots (Rf = 0.41, 0.35). The two components of the product mixture were separated by

flash chromatography and designated as 2.9u and 2.9l corresponding to Rf = 0.41, 0.35

respectively. Characterization of the two products by 1H-NMR and HRMS showed that

they were both stereoisomers of 2.9. Since there are three chiral centers in 2.9, there can

be four diastereomers, so each of the products isolated may still be a mixture of two or

more stereoisomers of product 2.9.

Hydrolysis of the methyl ester 2.9 was accomplished by treatment with sodium

hydroxide in water/methanol/THF (2/5/3, v/v/v), followed by acidification with

hydrochloric acid, as shown in Scheme 2.13. The 1H-NMR spectrum of the product

showed the disappearance of the methyl peak of the methyl ester at δ 3.6 – 3.7 ppm.

Since the final product iso[4]LGE2-d6 in an aldehyde form is not stable and is not

suitable for long term storage, the acetal 2.10 is useful as a precursor of iso[4]LGE2-d6 that is suitable for long term storage at -20 oC and that can be quickly and easily hydrolyzed to generate iso[4]LGE2-d6 as needed.

2.2.8 Generation of iso[4]LGE2-d6

Generation of iso[4]LGE2-d6 was accomplished by hydrolyzing the dimethyl

acetal 2.10 in dilute acetic acid for 3 h followed by evaporation of the solvent under

35 reduced pressure and high vacuum, as shown in Scheme 2.14. The yield was estimated by integration of the aldehyde hydrogen compared with the three hydrogens of the terminal methyl group of the lower side chain (Fig. 2.3), as in the earlier synthesis of unlabeled

4 iso[4]LGE2.

0.52 3.00

Fig 2.3 Estimation of the yield of hydrolysis of acetal 2.10 to generate iso[4]LGE2-d6

2.3 CONCLUSIONS

Synthesis of deuterium-labeled iso[4]LGE2-d6 was accomplished by an improved

procedure based on the previous synthesis of iso[4]LGE2. The new procedure eliminates

two bottlenecks from the original method. Deuterium was introduced in the top chain,

and THF-d8 was used as the source of deuterium. The top chain intermediate 2.3 for iso[4]LGE2-d6 was assembled through ring opening of THF-d8, alkylation of diethyl(2-

oxopropyl)phosphonate, deprotection of the TBDMS ether, oxidation of the alcohol and

esterification. The bottom side chain 2.6 was synthesized following the procedure in the

improved synthesis of iso[4]LGE2. After the top chain 2.3 was converted to enone 2.5 as a cis- and trans- mixture, the bottom side chain 2.6 was connected to the enone 2.5

through a Michael addition mechanism via a cyanocuprate reaction to produce the

iso[4]LGE2-d6 carbon skeleton 2.8, which after removal of the TBDMS ether protecting

group and hydrolysis of the methyl ester delivered the iso[4]LGE2-d6 precursor 2.10 as a

dimethyl acetal that is suitable for long term storage. The final product iso[4]LGE2-d6

2.11 is then readily produced by rapid hydrolysis of the precursor 2.10. The synthesis did not generate a pure diastereomer as in the previous synthesis. This is satisfactory since all diastereomers are produced through the free radical oxidation of arachidonic acid and it was not desirable to only generate one diastereomer.

2.4 EXPERIMENTAL

General Methods. Proton nuclear magnetic resonance (1H-NMR) spectra were

recorded on a Varian Unity Inova 400 NMR system with Oxford AS400 Actively

Shielding Magnet. Deuterium nuclear magnetic resonance (2H-NMR) spectra and carbon

nuclear magnetic resonance (13C-NMR) spectra were recorded on a Varian Inova AS600 spectrometer operating at 92 MHz and 151 MHz respectively. Hydrogen and deuterium chemical shifts are reported in parts per million (ppm) on the δ scale referenced to the

1 2 solvent CDCl3 (δ 7.26) and CHCl3 (δ 7.26) respectively. H/ H-NMR spectral data are tabulated in terms of multiplicity of hydrogen / deuterium absorption (s, singlet; d, doublet; dd, doublet of doublets; t, triplet; q, quartet; m, multiplet), coupling constants

(Hz) and number of protons / deuteriums. Carbon chemical shifts are reported in ppm on the δ scale referenced to the solvent CDCl3 (δ 77.0). The quintets of deuterated carbons are reported as the chemical shifts of the center peak in parenthesis. All high-resolution mass spectra were recorded on a Thermal Scientific LTQ FT Ultra mass spectrometer.

All solvents were distilled under a nitrogen atmosphere prior to use.

Tetrahydrofuran was distilled over sodium / benzophenone. Methylene chloride and acetonitrile were obtained dry from Acros Chemical Company. THF-d8 was purchased

from CDN ISOTOPES with 99.5 atom % D. 2,2-Dimethoxyglyoxal was purchased from

Lancaster Organics as a 60% aqueous solution. All other chemicals were obtained from

Aldrich or Acros (Fisher Scientific).

Chromatography was performed with ACS grade solvents. Thin-layer chromatography (TLC) was performed on glass plates precoated with silica gel

(Kieselgel 60 F254, E. Merck, Darmstadt, Germany). Rf values are quoted for plates of thickness 0.25 mm. The plates were visualized by exposure to iodine vapor or by heating the plates after dipping in a 20% solution of phosphomolybdic acid in ethanol. Flash

chromatography was performed on Purasil 60A silica gel(230 – 400 mesh) supplied by

Whatman.

tert-Butyl(1,1,2,2,3,3,4,4-d8-4-iodobutoxy)dimethylsilane (2.1)

To a flame-dried flask were added tert-butyldimethylsilyl chloride (23.6 g, 0.16

mmol), anhydrous acetonitrile (85 mL), tetrahydrofuran (THF)-d8 (9.68 g, 0.12 mmol and

sodium iodide (23.4 g, 0.16 mmol). The resulting turbid yellowish brown mixture was

stirred under argon at room temperature in the dark for 2 days. The solution was then

filtered through a sintered-glass funnel and the solid was washed with 25% ethyl acetate

in hexanes. The filtered clear wine red solution was then rinsed with 50% saturated

sodium thiosulfate (5 x 40 mL) in a separatory funnel to deliver a clear colorless solution.

The combined organic layer was washed with water and then brine, dried over anhydrous

magnesium sulfate, and concentrated under reduced pressure to afford a clear tan residue.

The crude product was purified by flash chromatography on silica gel with 100% hexanes

(Rf = 0.28) to deliver a slight turbid lightly yellow oil (22.15 g, 68.7 mmol, 58%).

1 H-NMR (400 MHz, CDCl3) δ 0.84 (s, 9H), 0.00 (s, 6H).

2 H-NMR (92 MHz, CHCl3) δ 3.58 (s, 2D), 3.18 (s, 2D), 1.84 (s, 2D), 1.54 (s, 2D).

13 C-NMR (151 MHz, CDCl3) δ (61.07), (32.29), (29.01), 25.91, 18.27, (6.51), -5.33.

+ + HRMS (EI) m/z 323.11378 (MH ) calcd for C10H16D8OISi found 323.11388; 191.01673

+ + (MH - HOSi(CH3)2C4H9) calcd for C4D8I found 191.01679.

Diethyl (4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)-2-oxoheptan-3-

yl)phosphonate (2.2)

To a solution of sodium hydride (380 mg of 60% dispersion in mineral oil, 9.5

mmol) in freshly distilled dry THF (5 mL) was added diethyl(2-oxopropyl)phosphonate

(1.85 mL, 9.2 mmol) in freshly distilled dry THF (5 mL) via cannula at -40 oC under

argon. The resulting turbid yellow solution was stirred at the temperature for 15 min and cold bath was then removed. After stirring for 2 h the solution became almost clear yellow. Then 2.1 (2.01 g, 6.2 mmol) in freshly distilled dry THF (4 mL) was added to the

solution at room temperature via cannula. The mixture, which was almost clear tan in color, was covered with aluminum foil and stirred under argon at room temperature.

After 10 days, the reaction was quenched by adding 12 mL of saturated NH4Cl, adjusted pH to about 5 by addition of 2 M HCl, and then extracted with ethyl acetate. The combined organic layer was rinsed with water, then brine, dried over anhydrous magnesium sulfate and concentrated by rotary evaporation to obtain a clear bright yellow

oil. TLC analysis (Rf = 0.28 in 100% hexanes) of the crude product showed that there

was unreacted starting material 2.1. The crude product was purified by flash chromatography using hexanes, 5%, 50% and 100% ethyl acetate in hexanes to obtain recovered 2.1 (1.18 g) and 2.2 (766.2 mg, 1.97 mmol, Rf = 0.5 in 100% ethyl acetate) in

77% yield based on the amount of 2.1 consumed.

1 H-NMR (400 MHz, CDCl3) δ 4.18 – 4.00 (4H), 3.09 (d, J = 24.4 Hz, 1H), 2.28 (s, 3H),

1.29 (tdd, J = 7.0, 1.4, 0.4 Hz, 6H), 0.85 (s, 9H), 0.00 (s, 6H).

2 H-NMR (92 MHz, CHCl3) δ 3.53 (s, 2D), 2.10 – 1.60 (2D), 1.44 (s, 1H), 1.26 (s, 2D).

13C-NMR (151 MHz, CDCl3) δ 204.05, 62.75, (61.96), 53.76 (d, J = 124.6 Hz, split by

31P), 31.36, 26.13, (25.59), (23.97), 18.51, 16.56, -5.11.

+ + HRMS (EI) m/z 389.27228 (MH ) calcd for C17H30D8O5PSi found 389.27219.

Diethyl (4,4,5,5,6,6,7,7-d8-7-hydroxy-2-oxoheptan-3-yl)phosphonate (2.3)

O O D2 D2 D2 D2 OH AcOH/H2O (3/1, v/v) C C C C OTBDMS C C C C 96% D D D D 2 2 2 2 O P OEt O P OEt OEt 2.3 OEt 2.2

The silyl ether 2.2 (1.43 g, 3.7 mmol) was dissolved in 12 mL of a freshly

prepared solution of acetic acid in water (3/1, v/v). The turbid light yellow solution was

stirred overnight at room temperature and became clear light tan in color. The solution

was concentrated by rotary evaporation to deliver a turbid light yellow liquid. The crude

product was purified by flash chromatography using 5% methanol in ethyl acetate (Rf =

0.23) to obtain 2.3 as a slightly turbid light yellow oil (0.98 g, 3.6 mmol, 96%).

1 H-NMR (400 MHz, CDCl3) δ 4.17 – 4.04 (4H), 3.12 (d, J = 24.6 Hz, 1H), 2.30 (s, 3H),

1.31 (t, J = 7.1 Hz, 6H).

2 H-NMR (92 MHz, CDCl3) δ 3.58 (s, 2D), 2.10 – 1.60 (2D), 1.49 (s, 2H), 1.29 (s, 2D).

13 31 C-NMR (151 MHz, CDCl3) δ 204.01, (61.34), 53.62 (d, J = 124.8 Hz, split by P),

31.41, (31.37), (25.36), (23.78), 16.53.

+ + HRMS (EI) m/z 275.18580 (MH ) calcd for C11H16D8O5P found 275.18577; 292.21235

+ + (M·NH4 ) calcd for C11H19D8NO5P found 292.21240.

Methyl 2,2,3,3,4,4-d6-5-(diethoxyphosphoryl)-6-oxoheptanoate (2.4)

To a solution of alcohol 2.3 (1.49 g, 5.4 mmol) in acetone (10 mL) was slowly

added Jones reagent31 (10 mL) at -10 oC. The resulting bright orange mixture was stirred

at -10 oC for 30 min until the dark red color persistent. The cold bath was then removed

and the solution was stirred for another 6 h. The reaction was quenched by slowly adding

isopropyl alcohol until the solution turned blue. The mixture was filtered through a short

celite pad, and concentrated by rotary evaporation to afford a clear yellow oil.

A mixture of 10 mL 50% aqueous KOH solution and 20 ml of ethyl ether was cooled to -5 ⁰C in a clean, nick-free Erlenmeyer flask. N-Nitrosomethylurea (2 g) was added in one portion and the resulting mixture was stirred magnetically for 10 min at the temperature. The cold bath was then removed and both layers rapidly turned yellow. The

Erlenmeyer flask was immersed in an acetone / dry ice bath to freeze the bottom aqueous layer and the upper diethyl ether layer containing diazomethane was decanted into another clean Erlenmeyer flask with KOH pellets as drying agent.

The crude oxidation product was transferred into a clean Erlenmeyer flask with 25 mL of ethyl ether and the resulting mixture was magnetically stirred. Freshly prepared diazomethane solution was then added until a bright yellow color persisted. The reaction was then quenched by addition of a few drops of acetic acid until the solution became colorless. The solution was concentrated by rotary evaporation to afford a turbid yellowish oil. The residue was purified by flash chromatography using 80% ethyl acetate in hexanes (Rf = 0.25) to obtain 2.4 as a clear light yellow oil (1.09 g, 3.6 mmol, 70%).

1 H-NMR (400 MHz, CDCl3) δ 4.15 – 4.02 (4H), 3.63 (s, 3H), 3.10 (d, J = 24.8 Hz, 1H),

2.30 (s, 3H), 1.29 (td, J = 7.1, 1.1 Hz, 6H).

2 H-NMR (92 MHz, CDCl3) δ 2.27 (s, 2D), 2.10 – 1.65 (2D), 1.53 (s, 2D).

13 C-NMR (151 MHz, CDCl3) δ 203.55, 173.53, 62.79, 53.43 (d, J = 124.8 Hz, split by

31P), 51.71, (32.94), 31.34, (25.13), (22.79), 16.49.

+ + HRMS (EI) m/z 301.16816 (MH ) calcd for C12H18D6O6P found 301.16811; 318.19471

+ + (M·NH4 ) calcd for C12H21D6NO6P found 318.19473.

Methyl 2,2,3,3,4,4-d6-5-acetyl-7,7-dimethoxyhept-5-enoate (2.5)

A suspension of sodium hydride (58.4 mg of 60% dispersion in mineral oil, 1.5 mmol) in freshly distilled dry THF(5 mL) was magnetically stirred at -20 oC. A solution

of 2.4 (310.6 mg, 1.04 mmol) in freshly distilled dry THF (2 mL) was cooled to -20 oC

and added dropwise via cannula. After stirring for 30 min the solution, which became

bright yellow, was warmed to -5 oC and stirring was continued for another 2 h and then

the mixture was cooled to -78 oC.

Meanwhile aqueous 2,2-dimethoxyglyoxal (60 w.t. %, 4.2 mL) was extracted

with diethyl ether. The organic layers were combined, dried over anhydrous magnesium

sulfate, and filtered through a celite pad under argon. This solution was cooled to -78 oC and added dropwise to the phosphonate solution via cannula. The resulting solution was stirred under argon at -78 oC and allowed to warm to room temperature overnight during

which it became colorless. The mixture was then cooled to -20 oC and quenched by

adding aqueous sodium dihydrogen phosphate solution (1 g in 10 mL water) and brine

(10 mL) and extracted with ethyl acetate. The organic layers were combined, washed

with water, then brine, and dried over anhydrous magnesium sulfate, filtered and

concentrated by rotary evaporation to afford a clear bright yellow oil. TLC analysis (25%

ethyl acetate in hexanes) showed two product spots (Rf = 0.26, 0.30) corresponding to the cis- and trans- isomers. The residue was purified by flash chromatography using 25%

ethyl acetate in hexanes, and the two isomers were collected together to obtain the

product 2.5 (249.9 mg, 1.0 mmol, 96%).

1 H-NMR (400 MHz, CDCl3) (mixture of cis- and trans- isomers) δ [6.44 (d, J = 6.4 Hz),

5.44 (d, J = 5.5 Hz) 1H], [5.16 (d, J = 6.4 Hz), 5.03 (d, J = 5.5 Hz), 1H], 3.65 (s, 3H),

[3.36 (s), 3.29 (s), 6H], [2.31 (s), 2.27 (s), 3H].

2 H-NMR (92 MHz, CDCl3) δ 2.45 – 2.10 (4D), 1.75 – 1.40 (2D).

13 C-NMR (151 MHz, CDCl3) (mixture of cis- and trans- isomers) δ 204.63, 199.51,

173.92, 145.41, 144.22, 138.06, 128.47, 100.03, 99.99, 53.22, 53.11, 51.72, 51.63,

(32.98), 30.09, 25.99, (24.74), (23.45).

+ + HRMS (EI) m/z 268.20256 (M·NH4 ) calcd for C12H18D6NO5 found 268.20262;

+ + 273.15796 (MNa ) calcd for C12H14D6O5Na found 273.15800.

(7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-9-(tert-butyldimethylsilyloxy)-6-

(dimethoxymethyl)heptadeca-7,11-dienoate (2.8)

Copper cyanide (198.5 mg, 2.2 mmol) in dry THF (5 mL, freshly distilled) in a

flame dried round bottom flask was cooled to 0 oC and methyl lithium (3.1 mL of 1.6 M

solution in ethyl ether, 4.8 mmol) was added. The solution first became clear bright

yellow and finally slightly tan as the addition proceeded. The mixture was stirred for 40

min at 0 oC and then the cold bath was removed. Vinylstannane 2.6 (1.4 g, 2.4 mmol) in

dry THF (5 mL, freshly distilled) was added drop wise via cannula at room temperature.

The solution became dark red in color. After stirring for 2 h under argon, the solution was cooled to -78 oC, and the enone 2.5 (500.3 mg, 2.0 mmol) in dry THF (6 mL, freshly

distilled) was added dropwise via cannula. The solution turned orange. After stirring at

-78 oC for 40 min and -40 oC for 40 min, the yellow reaction mixture was quenched by

adding saturated aqueous ammonium chloride and ammonium hydroxide (9:1, v/v, 10

mL). The resulting mixture was extracted with ethyl acetate. The combined organic layer

was washed with water, then brine, and dried over anhydrous magnesium sulfate, filtered

through a short silica gel column and concentrated by rotary evaporation to afford a clear

light yellow oil. TLC analysis (25% ethyl acetate in hexanes) of the crude product

showed two spots corresponding to product 2.8 (Rf = 0.57) and unreacted enone (Rf =

0.32). The residue was purified by flash chromatography using 25% ethyl acetate in

hexanes to obtain the product 2.8 as a diastereomeric mixture (524.5 mg, 0.98 mmol, 58% yield based on the amount of enone consumed).

1 H-NMR (400 MHz, CDCl3) (diastereomic mixture) δ 5.63 – 5.19 (4H), 4.24 – 3.99 (2H),

3.64 – 3.60 (3H), 3.31 – 3.14 (6H), 2.78 – 2.47 (2H), 2.16 – 2.08 (3H), 1.97 (q, J = 6.8

Hz, 2H), 1.36 – 1.15 (8H), 0.92 – 0.80 (12H), 0.06 – -0.03 (6H).

2 H-NMR (92 MHz, CDCl3) δ 2.49 – 1.97 (s, 2D), 1.80 – 1.06 (4D).

+ + HRMS (EI) m/z 550.44045 (M·NH4 ) calcd for C29H52D6NO6Si found 550.44045;

+ + 555.39585 (MNa ) calcd for C29H48D6O6SiNa found 555.39599.

(7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9-hydroxyheptadeca-

7,11-dienoate (2.9)

A solution of silyl ether 2.8 (102.9 mg, 0.19 mmol) in THF (10 mL) was cooled to

0 oC and then tetra-n-butylammonium floride solution (TBAF, 1.0 M in THF with 5%

water, 3 mL) was added. After stirring at 0 oC for 15 min, the cold bath was removed and the clear tan solution was stirred at room temperature overnight under argon during which it became clear brown. The reaction was cooled to 0 oC and quenched with water

(30 mL). The mixture was extracted with ethyl acetate. The combined organic layer was

washed with water, then brine, and dried over anhydrous magnesium sulfate, filtered

through a short silica gel column and concentrated by rotary evaporation to afford a clear

brown oil. TLC analysis (30% ethyl acetate in hexanes) of the crude product showed two

product spots (Rf = 0.41, 0.35) corresponding to two groups of diastereoisomers (2.9u

and 2.9l). The crude product was purified by flash chromatography using 30% ethyl

acetate in hexanes and the two groups of diastereoisomers were collected separately. The

overall yield (65.5 mg, 0.17 mmol, 90%) was calculated based on the total weight of the

two groups of diastereoisomers.

1 2.9u: H-NMR (400 MHz, CDCl3)δ 5.77 – 5.23 (4H), 4.30 – 4.04 (2H), 3.73 – 3.58 (3H),

3.37 – 3.18 (6H), 2.80 – 2.50 (2H), 2.39 – 2.17 (2H), 2.16 – 2.10 (3H), 2.02 (q, J = 7.1

Hz, 2H), 1.42 – 1.16 (6H), 0.86 (t, J = 6.8 Hz, 3H).

2 H-NMR (92 MHz, CDCl3): δ 2.27 (3D), 1.41 (3D).

+ + HRMS (EI) m/z 436.35398 (M·NH4 ) calcd for C23H38D6NO6 found 436.35395;

+ + 441.30937 (MNa ) calcd C23H34D6O6Na found 441.30948.

1 2.9l: H-NMR (400 MHz, CDCl3) δ 5.76 – 5.25 (4H), 4.34 – 4.01 (2H), 3.71 – 3.59 (3H),

3.44 – 3.15 (6H), 2.80 – 2.47 (2H), 2.41 – 2.17 (2H), 2.17 – 2.06 (3H), 2.00 (d, J = 6.4

Hz, 2H), 1.37 – 1.11 (6H), 0.85 (t, J = 6.7 Hz, 3H).

2 H-NMR (92 MHz, CDCl3): δ 2.74 – 1.91 (3D), 1.91 – 1.14 (3D).

+ + HRMS (EI) m/z 436.35398 (M·NH4 ) calcd for C23H38D6NO6 found 436.35393;

+ + 441.30937 (MNa ) calcd C23H34D6O6Na found 441.30947.

(7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9-hydroxyheptadeca-7,11-

dienoic acid (2.10)

To a solution of methyl ester 2.9 (65.5 mg, 0.16 mmol) in water/methanol/THF

(2/5/3, v/v/v, 10 mL) was added a solution of sodium hydroxide (16 mg/mL, 10 mL) in

water/methanol/THF (2/5/3, v/v/v) at room temperature. The mixture was then

magnetically stirred for 3 h and TLC analysis with 10% methanol in methylene chloride showed the disappearance of 2.9 (Rf = 0.65) and appearance of a product (2.10) (Rf =

0.29). The reaction mixture was carefully acidified to pH 4 by addition of 2 M HCl during which the color of the solution changed from light yellow to colorless. The mixture was extracted with ethyl acetate, and the combined organic layer was dried over anhydrous magnesium sulfate, filtered through a celite pad and concentrated by rotary evaporation to afford a clear brown oil. 1H-NMR showed that the crude product was

already pure enough and no need for further purification. The yield is quantitative (63.0

mg, 0.16 mmol).

1 H-NMR (400 MHz, CDCl3)δ 5.77 – 5.26 (4H), 4.28 – 4.02 (2H), 3.36 – 3.19 (6H), 2.81

– 2.52 (2H), 2.40 – 2.17 (2H), 2.17 – 2.09 (3H), 2.06 – 1.95 (2H), 1.38 – 1.14 (6H), 0.86

(t, J = 6.8 Hz, 3H).

2 H-NMR (92 MHz, CDCl3) δ 2.70 – 1.88 (S, 2D), 1.44 (s, 4D).

+ + HRMS (EI) m/z 422.33832 (M·NH4 ) calcd for C22H36D6NO6 found 422.33834.

(7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6-formyl-9-hydroxyheptadeca-7,11-dienoic acid

(iso[4]LGE2-d6, 2.11)

A solution of 2.10 (20.9 mg, 0.052 mmol) in acetic acid/water (3/1, v/v, 10 mL) was magnetically stirred and the process of the reaction was monitored by TLC. After 3 h TLC analysis (10% methanol in methylene chloride) showed the product 2.11 (Rf =

0.46) and disappearance of the acetal 2.10 (Rf = 0.58). The solution was concentrated by rotary evaporation as well as high vacuum. The purity of final product 2.11 was estimated by the ratio of integration of the aldehyde hydrogen (δ 9.52,dt, J = 31.0, 13.5 Hz, 1H) to the three hydrogens (δ 0.87, t, J = 6.9 Hz, 3H) of the end methyl group of the lower side chain.

+ + HRMS (EI) m/z 358.28590 ([M-H2O]·NH4 ) calcd for C20H28D6NO4 found 358.28595;

+ + + 359.26991 (MH ) calcd for C20H27D6O5 found 359.26995; 359.28925 ([M-H2O]·NH4 )

13 + + calcd for C19 CH28D6NO4 found 359.28931; 376.29646 (M·NH4 ) calcd for

+ C20H30D6NO5 found 376.29562.

2.5 REFERENCES

1. Charvet, C.; Liao, W. L.; Heo, G. Y.; Laird, J.; Salomon, R. G.; Turko, I. V.;

Pikuleva, I. A., Isolevuglandins and mitochondrial enzymes in the retina: Mass spectrometry detection of post-translational modification of sterol-metabolizing

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2. Li, W.; Laird, J. M.; Lu, L.; Roychowdhury, S.; Nagy, L. E.; Zhou, R.; Crabb, J.

W.; Salomon, R. G., Isolevuglandins covalently modify phosphatidylethanolamines in vivo: Detection and quantitative analysis of hydroxylactam adducts. Free Radic Biol Med

2009, 47, (11), 1539 - 1552.

3. Laird, J. M. Isolevuglandin-derived Hydroxylactams: Total synthesis and generation of an isoLG-modified lysine. Case Western Reserve University, Cleveland,

2007.

4. Subbanagounder, G.; Salomon, R. G.; Murthi, K. K.; Brame, C.; Roberts, L. J., II,

Total synthesis of iso[4]-levuglandin E2. J Org Chem 1997, 62, (22), 7658 - 7666.

5. Amarnath, V.; Amarnath, K.; Masterson, T.; Davies, S.; Roberts, L. J., II, A simplified synthesis of the diastereomers of levuglandin E2. Synth Commun 2005, 35, (3),

397 - 408.

6. Kobayashi, Y.; Shimazaki, T.; Taguchi, H.; Sato, F., Highly stereocontrolled total synthesis of leukotriene B4, 20-hydroxyleukotriene B4, leukotriene B3, and their analogs.

J Org Chem 1990, 55, (19), 5324 - 5335.

7. Wavrin, L.; Viala, J., Clean and efficient oxidation of homoallylic and homopropargylic alcohols to β,γ-unsaturated aldehydes by the Dess-Martin periodinane.

Synthesis 2002, (3), 326 - 330.

8. Treilhou, M.; Fauve, A.; Pougny, J. R.; Prome, J. C.; Veschambre, H., Use of

biological catalysts for the preparation of chiral molecules. 8. Preparation of propargylic alcohols. Application in the total synthesis of leukotriene B4. J Org Chem 1992, 57, (11),

3203 - 3208.

9. Ireland, R. E.; Liu, L., An improved procedure for the preparation of the Dess-

Martin periodinane. J Org Chem 1993, 58, (10), 2899 - 2899.

10. Frigerio, M.; Santagostino, M.; Sputore, S., A user-friendly entry to 2- iodoxybenzoic acid (IBX). J Org Chem 1999, 64, (12), 4537 - 4538.

11. Dess, D. B.; Martin, J. C., A useful 12-I-5 triacetoxyperiodinane (the Dess-Martin

periodinane) for the selective oxidation of primary or secondary alcohols and a variety of

related 12-I-5 species. J Am Chem Soc 1991, 113, (19), 7277 - 7287.

12. Renaldo, A. F.; Labadie, J. W.; Stille, J. K., Palladium-catalyzed coupling of acid chlorides with organotin reagents: Ethyl (E)-4-(4-nitrophenyl)-4-oxo-2-butenoate. Org

Synth 1989, 67, 86 - 97.

13. Layer, R. W., Reaction of epoxides with 2,6-di-tert-butylphenol. J Org Chem

1981, 46, (25), 5224 - 5225.

14. Bencivengo, D.; San Filippo, J., Jr., Nonprotic procedure for transesterification of methyl esters. J Org Chem 1981, 46, (25), 5222 - 5224.

15. Keck, G. E.; Heumann, S. A., Diasteroselective synthesis of cyclopentapyridazinones via radical cyclization: Synthetic studies toward halichlorine.

Org Lett 2008, 10, (21), 4783 - 4786.

16. Salomon, R. G.; Miller, D. B.; Raychaudhuri, S. R.; Avasthi, K. L., K.; Levison,

B. S., Prostaglandin endoperoxides. 15. Asymmetric total synthesis of levuglandin E2. J

Am Chem Soc 1984, 106, (26), 8296 - 8298.

17. Miller, D. B.; Raychaudhuri, S. R.; Avasthi, K.; Lal, K.; Levison, B.; Salomon, R.

G., Prostaglandin endoperoxides. 25. Levuglandin E2: Enantiocontrolled total synthesis of a biologically active rearrangement product from the prostaglandin endoperoxide

PGH2. J Org Chem 1990, 55, (10), 3164 - 3175.

18. von Pechmann, H., Ueber diazomethan. Ber 1894, 27, 1888 - 1891.

19. Barkawi, L. S.; Cohen, J. D., A method for concurrent diazomethane synthesis and substrate methylation in a 96-sample format. Nat Protoc 2010,5, (10), 1619 - 1626.

20. Borzsonyi, M.; Sajgo, K.; Torok, G.; Pinter, A.; Tamas, J.; Kolar, G.;

Spiegelhalder, B., Transnitrosating activity of N-nitroso-N-methyl-p-toluenesulfonamide in rats and human gastric juice. Neoplasma 1988, 35, (3), 257 - 262.

21. Morandi, B.; Carreira, E. M., Iron-catalyzed cyclopropanation in 6 M KOH with in situ generation of diazomethane. Science 335, (6075), 1471 - 1474.

22. Yamamoto, M.; Ishiwata, H.; Yamada, T.; Tanimura, A.; Tomita, I., HPLC determination of N-nitrosomethylurea in biological fluids by reaction with 4- nitrothiophenol. Food Chem Toxicol 1986, 24, (3), 247 - 250.

23. Arndt, F., Diazomethan. Org Synth, Coll Vol 1943, 2, p.165.

24. Taylor, R. J. K., Organocopper reagents: A practical approach. Oxford

University Press: Oxford, 1994.

25. Lipshutz, B. H.; Wilhelm, R. S.; Kozlowski, J. A., The chemistry of higher order organocuprates. Tetrahedron 1984, 40, (24), 5005 - 5038.

26. Lipshutz, B. H.; Sengupta, S., Organocoper Reagents: Substitution, conjutate

addition, carbo/metallocupration and other reactions. Org Reactions (New York) 1992, 41,

135 - 631.

27. Lipshutz, B. H., Recent progress in higher order cyanocuprate chemistry.

AdvMetal-Organic Chem 1995, 4, 1 - 64.

28. Lipshutz, B. H., The evolution of higher order cyanocuprates. Synlett 1990, (3),

119 - 128.

29. Lipshutz, B. H., Applications of higher-order mixed organocuprates to organic

synthesis. Synthesis 1987, (4), 325 - 341.

30. Behling, J. R.; Babiak, K. A.; Ng, J. S.; Campbell, A. L. M., R.; Koerner, M.;

Lipshutz, B. H., In-situ cuprate formation via transmetalation between vinylstannanes and

higher order cyanocuprates. J Am Chem Soc 1988, 110, (8), 2641 - 2643.

31. Shriner, R. L.; Hermann, C. K. F.; Morrill, T. C.; Curtin, D. Y.; Fuson, R. C., The systematic identification of organic compounds. 8th ed.; Wiley: 2004.

Chapter 3

Total Synthesis of Deuterium-labeled isoLevuglandin E2-d6

3.1 BACKGROUND

IsoLGE2 is one of the structural isomers of isolevuglandins generated from free radical-induced autoxidation of AA. Unlike the iso[n]LGs, which can only be generated through a free radical pathway, LGE2, one steroisomer of isoLGE2 can also be generated

through an enzymatic pathway. Although various stereoisomers of LGE2 were

successfully synthesized before,1 the reported methods2-4 for the synthesis of the top side

chain are not applicable for the synthesis of deuterium-labeled isoLGE2-d6 due to harsh

reaction conditions and the expense and difficulty of introducing deuterium. In this

chapter we described an efficient route developed to use readily available starting

materials (THF-d8 and propargyl alcohol). This method is also applicable for synthesis of

non-deuterated isoLGE2 from THF.

3.2 RESULTS AND DISCUSSION

Due to nonregioselective abstraction of allylic hydrogen, a cis double bond resides in the bottom chain in iso[4]LGE2 while a cis double bond is located in the top

chain in isoLGE2. The overall synthetic strategy for isoLGE2-d6 is very similar to that of

iso[4]LGE2-d6. First the top side chain, an allylic bromide, and the bottom side chain, a vinylstannane, are constructed respectively. Then the top side chain is reacted with diethyl(2-oxopropyl)phosphonate to generate a phosphonated ester, which is further converted to an enone as the Michael acceptor to which the vinylstannane side chain is connected to form the carbon skeleton of isoLGE2. After removal of the protecting

groups on both side chains and hydrolysis of the acetal, isoLGE2 is finally obtained.

Previously in the Salomon group, different stereoisomers of LGE2 were

successfully synthesized.1 In these syntheses, the top segment, cis-methyl 7-bromohept-5-

enoate, was synthesized via a literature method2, as shown in Scheme 3.1A. This 7-step

synthesis started with an inexpensive tetrahydropyran (THP) protected propargyl alcohol,

which is commercially available or can be readily prepared from even less expensive

propargyl alcohol.5-7 However, it included prolonged heating under reflux which requires

condensation devices, thus increasing the chance of lab accident, especially when left

unattended overnight.

Another reported method3, 4 starts with hex-5-ynoic acid and is accomplished in

four steps, as shown in Scheme 3.1B. In order to use a readily available and inexpensive

starting material for the synthesis of iso[4]LGE2-d6, deuterium was introduced into the

57 top side chain, thus this reported method is not applicable for the synthesis of isoLGE2-d6 since deuterated hex-5-ynoic acid is not commercially available.

Our synthesis, which starts with very inexpensive propargyl alcohol and the ring opening product 2.1 from THF-d8, is accomplished efficiently in five steps (Scheme 3.2), two steps shorter than the original synthesis of the unlabeled analog of the carboxylic

allylic bromide side chain intermediate.1 This method not only is applicable for synthesis of deuterated isoLGE2-d6. For unlabeled isoLGE2, although it is one step more than the

reported method shown in Scheme 3.1B, the synthesis can start with unlabeled THF, thus

makes this method even more affordable and attractive.

Top side chain 3.5 was reacted with diethyl(2-oxopropyl)phosphonate to generate

a phosphonate ester, which was further converted to an enone as the Michael acceptor to

which the vinylstannane side chain was connected to form the carbon skeleton of isoLGE2-d6. After removal of the protecting groups on both side chains and hydrolysis of

the acetal, isoLGE2-d6 was finally obtained.

3.2.1 Alkylation of propargyl alcohol

The strategy for alkylation of propargyl alcohol is to use two equivalents of base to deprotonate the alcohol as well as the terminal alkyne carbon to form a dianion and,

since the alkynyl anion is more neucleophilic than the oxygen anion, it is more favored to

react with the iodobutyl TBDMS ether 2.1 via an SN2 mechanism to accomplish the

alkylation at the alkyne terminal carbon. The alcohol is recovered after workup to

produce 3.1.

Several bases have been reported8-11 to be useful for alkylation of propargyl

alcohol. Commercially available LiNH2 did not work for this reaction at all. Li /NH3 (l.) worked but with a poor yield. One reason the Li /NH3 (l.) system was abandoned is that it

is hard to control the amount of lithium amide generated due to the difficulty of

accurately measuring the weight of lithium wire added into the liquid ammonia. Excess

base might destroy the alkyl iodide. This might be the reason for the poor yield.

Furthermore, the use of poisonous ammonia and the complexity of reaction apparatus are

other drawbacks of this approach.

The best yield was achieved by using nBuLi as base. When optimizing the

reaction conditions, the ratio of alcohol to iodoalkane was found to play an important role in the yield. Since the deuteratued iodoalkane 2.1 is more valuable than propargyl alcohol, in order to consume all of the deuteratued iodoalkane 2.1, two equivalents of propargyl

alcohol was always used while changing other conditions, such as the reaction time, the

60 amount of HMPA, etc, but the yield was always around 30%. Later when 1 equivalent of propargyl alcohol was used, the best yield (61%) was finally delivered. Further decrease in the amount of propargyl alcohol did not result in a better yield. The reason why an excess of propargyl alcohol gave bad yield might be the excess dianion generated from propargyl alcohol which might have caused some side reactions, such as elimination or hydrolysis of the silyl ether.

Base R-OH:2.1 Yield

LiNH2 n/a 0

Li/NH3 (l) n/a poor 2:1 ca 30% nBuLi 1:1 61%

Table 2.1 The yield of alkylation of propargyl alcohol with 2.1 using different bases

The use of hexamethylphosphoramide (HMPA) is essential for this reaction.

HMPA is commonly used as a solvent for organometallic compounds. The P-O bond is highly polarized, with significant negative charge residing on the oxygen atom. This allows it to accelerate some SN2 reactions by selectively solvating cations.

Examination of the 1H-NMR spectrum of the product showed a singlet at δ 4.20 ppm corresponding to the methylene protons α to the hydroxyl group.

3.2.2 Acetylation of alkylated propargyl alcohol (3.1)

The propargyl alcohol of 3.1 was protected as an acetate to prevent its oxidation in the next step. The oxidation was to be done under acidic conditions using Jones reagent. THP cannot be used as a protecting group since it is acid sensitive, otherwise the synthesis could have used THP protected propargyl alcohol to save a step.

Acetylation was accomplished by adding three equivalents of acetic anhydride to a solution of 3.1 in pyridine at low temperature followed by stirring at room temperature for an additional 2 h. The excess of acetic anhydride was quenched by addition of ethanol.

The crude product was purified by flash chromatography to deliver product 3.2 with a yield of 96%. Confirmation of acetylation is evident by 1H-NMR spectroscopy that

shows a singlet at 2.05 ppm corresponding to the methyl of the acetyl group. Also the

proton resonance of the methylene α to the hydroxyl group on 3.1 shifted downfield from

4.20 ppm to 4.61 ppm on 3.2 due to more shielding by the newly formed acetyl group.

3.2.3 Generation of 3.3: deprotection, oxidationand esterification of 3.2

Removal of the TBDMS protecting group and oxidation of the alcohol to a

carboxylic acid were accomplished in one step using Jones reagent via established

procedures.12

Conversion of the intermediate acetoxy acid to the hydroxy acid was

accomplished simply by extraction of the crude product from previous step with 10%

aqueous NaOH solution, followed by acidification with 6 M HCl.10 The crude product

was used directly for esterification using freshly prepared diazomethane following the

procedure described in Chapter 2 to finally obtain 3.3. After purification by flash

chromatography an overall yield of 32% was obtained for the above four steps.

3.2.4 Selective hydrogenation of the triple bond to a cis double bond using P-2 nickel

boride

Although both the previously reported methods used Lindlar’s catalyst to effect the hydrogenation, we used inexpensive, easily prepared P-2 nickel boride as the catalyst,3, 13-17 which can convert alkynes stereoselectively to cis-alkenes in very good

yield (89%). P-2 nickel, a nearly colloidal black suspension, was prepared by treating a vigorously stirred solution of nickel acetate in 95% ethanol with ethanolic sodium

63 borohydride under hydrogen. In the presence of ethylenediamine the nearly pure cis isomer is obtained.

3.2.5 The Appel reaction: bromination of 3.4 to generate the top side chain precurosor 3.5

Bromination of the allylic alcohol 3.4 was achieved by established procedures18, 19 to finally obtain the deuterated top side chain precursor 3.5 for isoLGE2-d6, as shown in

Scheme 3.7.

3.2.6 Generation of enone-d6 3.7

O O O D2 D2 D2 D2 C C C C a b C COOCH3 C COOCH3 D 71% D 83% 2 2 H3CO O P OEt O P OEt

OCH3 3.7 OEt OEt 3.6

a. NaH, THF, then 3.5; b. NaH, THF, 2,2-dimethoxyglyoxal

Scheme 3.8 Generation of enone-d6 3.7

After diethyl(2-oxopropyl)phosphonate was alkylated with the top side chain precursor 3.5 (Scheme 3.8), the product 3.6 was converted to the Michael acceptor 3.7

64 following the same strategy as the synthesis of iso[4]LGE2-d6 discussed in Chapter 2.

Unlike the enone-d6 2.5, which clearly showed two spots on the TLC plate corresponding to two isomers, the enone-d6 3.7 just showed one spot, and the presumed mixture of isomers was collected together after flash chromatography.

3.2.7 Elaboration of isoLGE2-d6 (3.12) from enone-d6 3.7

The bottom side chain 3.8 was synthesized via the reported method,20 except that we did not do enzymatic resolution to get a pure enantiomer since isoLGs are, by definition, already mixtures of enantiomers via radical formation.

The connection of the bottom side chain 3.8 to enone-d6 3.7 used the similar

cyanocuprate chemistry as did in the synthesis of iso[4]LGE2-d6 discussed in Chapter 2 to

afford the isoLGE2-d6 carbon skeleton 3.9. Following removal of the TBDMS group and

then hydrolysis of the methyl ester, isoLGE2-d6 acetal 3.11 was finally obtained. It is

o used as a precursor of isoLGE2-d6 that is suitable for long term storage at -20 C. From a

rapid hydrolysis of the acetal 3.11 the final product isoLGE2-d6 (3.12) was obtained. The yield was estimated by the integration of the aldehyde hydrogen compared with the three

hydrogens of the terminal methyl group of the lower side chain, as did in the iso[4]LGE2- d6 synthesis.

3.3 CONCLUSIONS

A synthesis of deuterated isoLGE2-d6 was successfully developed. By using

readily available starting materials (THF-d8 and propargyl alcohol) this method is also

applicable for synthesis of non-deuterated isoLGE2 from THF. It is noteworthy that ring

opening product 2.1 from THF-d8 can be used for synthesis of both iso[4]LGE2-d6 and

isoLGE2-d6. Use of non-THP-protected propargyl alcohol reacting with two equiv nBuLi

saved the step of THP deprotection. A one pot reaction for converting propargyl acetate

3.2 to methyl hydroxylalkynoate 3.3 avoided tedious purification of intermediates. By following a similar strategy as in the synthesis of iso[4]LGE2-d6, the isoLGE2-d6 carbon skeleton 3.9 was constructed. After the removal of protecting groups on the side chains, a stable dimethoxyacetal precursor 3.11 was obtained that is suitable for long time storage

o at -20 C. It can be quickly and easily hydrolyzed to produce the final product isoLGE2-d6

(3.12). As in the previous synthesis of iso[4]LGE2-d6,the synthesis did not generate a

pure diastereomer.

3.4 EXPERIMENTAL

General Methods. Proton nuclear magnetic resonance (1H-NMR) spectra were

recorded on a Varian Unity Inova 400 NMR system with Oxford AS400 Actively

Shielding Magnet. Deuterium nuclear magnetic resonance (2H-NMR) spectra and carbon

1 2 solvent CDCl3 (δ = 7.26) and CHCl3 (δ 7.26) respectively. H/ H-NMR spectral data are

tabulated in terms of multiplicity of hydrogen / deuterium absorption (s, singlet; d,

doublet; dd, doublet of doublets; t, triplet; q, quartet; m, multiplet), coupling constants

(Hz) and number of protons / deuteriums. Carbon chemical shifts are reported in ppm on

the δ scale referenced to the solvent CDCl3 (δ=77.0). The quintets of deuterated carbons are reported as the chemical shifts of the center peak in parenthesis. All high-resolution mass spectra were recorded on a Thermal Scientific LTQ FT Ultra mass spectrometer.

All solvents were distilled under nitrogen atmosphere prior to use.

Tetrahydrofuran was distilled over sodium / benzophenone. Methylene chloride and acetonitrile were obtained dry from Acros Chemical Company. THF-d8 was purchased from CDN ISOTOPES with 99.5 atom % D. 2,2-Dimethoxyglyoxal was purchased from

Lancaster Organics as a 60% aqueous solution. All other chemicals were obtained from

Sigma Aldrich or Acros (Fisher Scientific).

Chromatography was performed with ACS grade solvents. Thin-layer chromatography (TLC) was performed on glass plates precoated with silica gel

(Kieselgel 60 F254, E. Merck, Darmstadt, Germany). Rf values are quoated for plates of thickness 0.25 mm. The plates were visualized by exposure to iodine vapor or by heating the plates after dipping in 20% solution of phosphomolybdic acid in ethanol. Flash chromatography was performed on Purasil 60A silica gel(230 – 400 mesh) supplied by

Whatman.

4,4,5,5,6,6,7,7-d8-7-((tert-Butyldimethylsilyl)oxy)hept-2-yn-1-ol (3.1)

To a flamed-dried flask equipped with a magnetic stirring bar was added

propargyl alcohol (99%, 367 mL, 6.3 mmol), hexamethylphosphoramide (HMPA, 5 mL)

and freshly distilled dry THF (5 mL). The solution was cooled to -40 oC and n-

butyllithium (1.6 M in hexanes, 7.7 mL, 2 equiv) was added. The mixture was stirred for

1 h at -40 oC under argon. Iodide 2.1 (2.04g, 6.3 mmol) in freshly distilled dry THF (8

mL) was cooled to -40 oC added dropwise via cannula. The resulting mixture was stirred

under argon and allowed to slowly warm to room temperature. After 4 h, the mixture,

which was almost clear light yellow, was quenched by addition of 20 mL saturated

NH4Cl, extracted with diethyl ether, washed with water, then brine, and dried over

anhydrous magnesium sulfate and concentrated by rotary evaporation to deliver a clear

yellow oil. The residue was purified by flash chromatography using 10% ethyl acetate in

hexanes (Rf = 0.16) to obtain 3.1 (940.0 mg, 60%) as a clear light yellow oil.

1 H-NMR (400 MHz, CDCl3) δ 4.20 (s, 1H), 0.84 (s, 9H), 0.00 (s, 6H).

2 H-NMR (92 MHz, CHCl3) δ 3.58 (s, 2D), 2.18 (s, 2D), 1.65 – 1.40 (4D).

13 C-NMR (151 MHz, CDCl3) δ 86.12, 78.52, (61.78), 51.21, (30.60), 25.90, (23.81),

18.29, (17.70), -5.34.

+ HRMS (EI) m/z 251.22770 (MH ) calcd for C13H19D8O2Si found 251.22777; 273.20964

+ (MNa ) calcd for C13H18D8O2SiNa found 273.20964.

4,4,5,5,6,6,7,7-d8-7-((tert-Butyldimethylsilyl)oxy)hept-2-yn-1-yl acetate (3.2)

A solution of 3.1 (2.48 g, 9.9 mmol) in 15 mL pyridine was cooled to 0 oC and

acetic anhydride (2.8 mL, 29.7 mmol, 3 equiv) was slowly added. The mixture was

stirred for 40 min at the temperature under argon, and then allowed to warm to room temperature. After 2 h the mixture (clear light yellow) was quenched by addition ofethanol (5 mL) and concentrated by rotary evaporation to afford a clear yellow oil. The

residue was purified by flash chromatography using 5% ethyl acetate in hexanes (Rf =

0.25) to obtain 3.2 (2.79 g, 9.5 mmol, 96%).

1 H-NMR (400 MHz, CDCl3) δ 4.61 (s, 2H), 2.05 (s, 3H), 0.84 (s, 9H), 0.00 (s, 6H).

2 H-NMR (92 MHz, CDCl3) δ 3.58 (s, 2D), 2.19 (s, 2D), 1.52 (d,4D).

13 C-NMR (151 MHz, CHCl3) δ 170.26, 87.39, 74.01, (61.68), 52.78, (30.60),

25.89,(23.65), 20.75, 18.26, (17.71), -5.35.

+ HRMS (EI) m/z 315.22021 (MNa ) calcd for C15H20D8O3SiNa found 315.22024.

Methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-ynoate (3.3)

To a solution of 3.2 (5.2 g, 17.8 mmol) in acetone (30 mL) was slowly added

Jones reagent21 (2.5 M, 21.5 mL, 53.4 mmol, 3 equiv) at 0 oC. The mixture was

magnetically stirred at the temperature for 15 min. The cold bath was removed and the

mixture allowed to slowly warm to room temperature. After stirring overnight, the

reaction was quenched with 40 mL isopropyl alcohol, extracted with diethyl ether (5 x 50

mL), and washed with water, then brine, and dried over anhydrous magnesium sulfate,

and then concentrated by rotary evaporation to obtain the crude acetoxy acid as a clear

yellow oil (4.2 g, Rf = 0.4 in 50% ethyl acetate in hexanes). This crude product was

dissolved in ethyl acetate (20 mL) and extracted with 10% NaOH (4 x 20 mL). The

aqueous layers were collected and acidified with concentrated HCl to pH=1, followed by

extraction with ethyl acetate (5 x 20 mL). The organic layers were combined, washed

with water, then brine, and dried over anhydrous magnesium sulfate, filtered and

concentrated by rotary evaporation to afford the crude product, a hydroxy acid, as a clear

greenish oil (1.8 g, Rf = 0.17 in 50% ethyl acetate in hexanes).

The crude hydroxy acid was transferred into a 50 mL Erlenmeyer flask with

diethyl ether (20 mL) and stirred. A solution of freshly made diazomethane in diethyl

ether was added dropwise until a bright yellow color persisted, then excess diazomethane

was quenched by addition of a few drops of glacial acetic acid until the solution became colorless. The solution was concentrated by rotary evaporation to afford a turbid yellow oil. The residue was purified by flash chromatography using 30% ethyl acetate (Rf = 0.28) to obtain the product 3.3 (932.7 mg, 5.8 mmol, 32% for above 3 steps).

1 H-NMR (400 MHz, CDCl3) δ 4.22 (s, 2H), 3.66 (s, 3H).

2 H-NMR (92 MHz, CHCl3) δ 2.40 (s, 2D), 2.24 (s, 2D), 1.78 (s, 2D).

13 C-NMR (151 MHz, CDCl3) δ 173.77,84.71, 79.36, 51.57, 51.05, (31.96), (22.67),

(17.34).

+ + HRMS (EI) m/z 185.10553 (MNa ) calcd for C8H6D6O3Na found 185.10550.

(Z)-methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-enoate (3.4)

A solution of NaBH4 (612 mg, 15.9 mmol) in ethanol (50 mL) was added to a

solution of Ni(OAc)2·H2O (3.7 g, 15 mmol) in 95% ethanol (50 mL) under H2. The

resulting black suspension of nickel boride was stirred for 30 min, followed by addition

of ethylenediammine (2 mL). After 15 min a solution of 3.3 (502.7 mg, 3.1 mmol) in

ethanol (5 mL) was added, and the solution stirred for 4 h and was followed by TLC. The reaction was quenched by addition of 50 mL diethyl ether, filtered through a short silica gel column, and concentrated by rotary evaporation to afford a yellow oil. Note that the product cannot be dried under high-vacuum as it will evaporate. The residue was purified by flash chromatography using 40% ethyl acetate in hexanes (Rf = 0.28) to obtain 3.4

(454 mg, 2.8 mmol, 89%).

1 H-NMR (400 MHz, CDCl3) δ 5.65 (dt, J = 10.9, 6.9 Hz, 1H), 5.46 (d, J = 11.0 Hz, 1H),

4.14 (dd, J = 6.9, 1.3 Hz, 2H), 3.65 (s, 3H).

2 H-NMR (92 MHz, CHCl3) δ 2.27 (s, 2D), 2.08 (s, 2D), 1.66 (s, 2D).

13 C-NMR (151 MHz, CDCl3) δ 174.13, 131.31, 129.68, 58.25, 51.52, (32.42),

(25.58),(23.54).

+ + HRMS (EI) m/z 187.12118 (MNa ) calcd for C8H8D6O3Na found 187.12115.

(Z)-methyl 2,2,3,3,4,4-d6-7-bromohept-5-enoate (3.5)

To a solution of carbon tetrabromide (3.65 g, 11.0 mmol) in dry methylene

chloride (50 mL) was added a solution of 3.4 (903.2 mg, 5.5 mmol) in dry methylene

chloride (10 mL) and triphenylphosphine (2.88 g, 11.0 mmol). The resulting clear orange

red solution was stirred for 1.5 h under argon. The solvent was then removed by rotary

evaporation and the residue was purified by flash chromatography using 10% ethyl

acetate in hexanes (Rf = 0.38) to obtain the compound 3.5 (1.03 g, 4.5 mmol, 83%).

1 H-NMR (400 MHz, CDCl3) δ 5.74 (tt, J = 9.8, 7.2 Hz, 1H), 5.54 (d, J = 10.6 Hz, 1H),

4.00 – 3.87 (2H), 3.69 – 3.62 (3H).

2 H-NMR (92 MHz, CHCl3) δ 2.27 (d, J = 2.6 Hz, 2D), 2.09 (d, J = 7.7 Hz, 2D), 1.67 (s,

2D).

+ + HRMS (EI) m/z 249.03677 (MNa ) calcd for C8H7D6O2BrNa found 249.03682.

(Z)-methyl 2,2,3,3,4,4-d6-8-(diethoxyphosphoryl)-9-oxodec-5-enoate (3.6)

Toa solution of sodium hydride (338.1 mg of 60% dispersion in mineral oil, 8.5

mmol) in freshly distilled THF (20 mL) was added diethyl(2-oxopropyl)phosphonate (1.6

mL, 7.9 mmol) at room temperature under argon. The solution was stirred for 2 h, then

3.6 (1.09 g, 4.8 mmol) was added in freshly distilled dry THF (10 mL) via cannula. The

mixture was stirred overnight under argon, and then quenched with 10 mL saturated

NH4Cl solution. 2 M HCl was used to adjust the pH of the solution to around 5. The

solution was extracted with ethyl acetate (5 x 10 mL). The organic layers were combined

and washed with water, then brine, and dried over anhydrous magnesium sulfate and

concentrated by rotary evaporation to obtain a clear yellow liquid (2.05 g), which was

purified by flash chromatography using 80% ethyl acetate in hexanes (Rf = 0.3) to obtain

the product 3.6 (1.15 g, 3.4 mmol, 71%).

1 H-NMR (400 MHz, CDCl3) δ 5.47 – 5.18 (2H), 4.18 – 4.01 (4H), 3.66 – 3.62 (3H), 3.24

– 3.07 (m, 1H), 2.79 – 2.59 (m, 1H), 2.51 – 2.37 (m, 1H), 2.28 – 2.25 (3H), 1.34 – 1.27

(m, 6H).

2 H-NMR (92 MHz, CHCl3) δ 2.24 (d, J = 4.6 Hz, 2D), 1.99 (d, J = 10.6 Hz, 2D), 1.62 (s,

2D).

+ + HRMS (EI) m/z 341.19946 (MH ) calcd for C15H22D6O6P found 341.19943; 358.22601

+ + + (M·NH4 ) calcd for C15H25D6NO6P found 358.22605; 363.18141 (MNa ) calcd for

+ C15H21D6O6PNa found 363.18155.

(5Z)-methyl 2,2,3,3,4,4-d6-8-acetyl-10,10-dimethoxydeca-5,8-dienoate (3.7)

A suspension of sodium hydride (66.7 mg of 60% dispersion in mineral oil, 1.67 mmol) in freshly distilled THF (6 mL) was magnetically stirred at -20 oC. A solution of

3.6 (433.0 mg, 1.27 mmol) in freshly distilled THF (6 mL) was cooled to -20 oC and

added dropwise via cannula. The resulting solution was turbid light yellow. After stirring

for 20 min the solution was warmed to -5 oC and stirring continued for another 2 h and

then the mixture was cooled to -78 oC.

Meanwhile aqueous 2,2-dimethoxyglyoxal (60 w.t. %, 8 mL) was extracted with

diethyl ether. The organic layers were combined, dried over anhydrous magnesium

sulfate, filtered through a celite pad into a flame-dried round bottom flask under argon.

This solution was cooled to -78 oC and added dropwise to the phosphonate solution via

cannula.

The resulting solution was stirred under argon at -78 oC and then allowed to warm

to room temperature overnight. The turbid, colorless mixture was cooled to -20 oC and

quenched by adding aqueous sodium dihydrogen phosphate solution (1 g in 10 mL water)

and brine (10 mL) and extracted with ethyl acetate. The organic layers were combined,

washed with water, then brine, and dried over anhydrous magnesium sulfate, filtered and

concentrated by rotary evaporation to afford a clear bright yellow oil. The residue was

purified by flash chromatography using 20% ethyl acetate in hexanes (Rf = 0.2) to obtain

the product 3.7(343.1 mg, 1.18 mmol, 93%, mixture of two isomers).

1 H-NMR (400 MHz, CDCl3) δ 6.43 (t, J = 6.6 Hz, 1H), 5.37 – 5.30 (m, 1H), 5.23 – 5.11

(2H), 3.67 – 3.62 (3H), 3.34 (s,6H), 3.11 – 3.03 (2H), 2.32 – 2.30 (3H).

2 H-NMR (92 MHz, CHCl3) δ 2.25 (2D), 2.14 (2D), 1.63 (2D).

+ + HRMS (EI) m/z 308.23386 (M·NH4 ) calcd for C15H22D6NO5 found 308.23386.

(5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12-((tert-butyldimethylsilyl)oxy)-9-

(dimethoxymethyl)heptadeca-5,10-dienoate (3.9)

A 50 mL round bottom flask was flame dried and copper cyanide (362.5 mg, 4.05

mmol) added immediately under argon. Freshly distilled dry THF (5 mL) was added after

the flask cooled to room temperature. The flask was cooled to 0 oC and methyl lithium

(5.6 mL of 1.6 M solution in ethyl ether, 8.9 mmol, 2.2 equiv) was added. The solution

first became clear bright yellow and finally clear light tan as the addition proceeded. The

mixture was stirred for 40 min at 0 oC and then the cold bath was removed.

Vinylstannane side chain 3.8 (2.58 g, 4.86 mmol, 1.2 equiv) in dry THF (5 mL, freshly

distilled) was added dropwise via cannula at room temperature. After stirring for 2 h under argon, the solution was cooled to -78 oC, and the enone 3.7 (534.4 mg, 1.84 mmol)

in THF (5 mL, freshly distilled) was added dropwise via cannula. After stirring at -78 oC for 40 min and -40oC for 40 min, the reaction was quenched by adding saturated aqueous

ammonium chloride and ammonium hydroxide (9:1, v/v, 12 mL). The resulting mixture

was extracted with ethyl acetate. The combined organic layer was washed with water,

then brine, and dried over anhydrous magnesium sulfate, filtered through a short silica

gel column and concentrated by rotary evaporation to afford a clear light yellow oil. TLC

analysis (15% ethyl acetate in hexanes) of the crude product showed two spots of product

2.8 (Rf = 0.33, 0.25) and one spot of unreacted enone (3.7) (Rf = 0.15). The residue was

purified by flash chromatography using 15% ethyl acetate in hexanes. Two product spots,

designated as 3.9u and 3.9l corresponding to the spots Rf = 0.33, 0.25 respectively, were

collected separately and unreacted enone (3.7) was recovered. The overall yield is 68%

based on the amount of enone (3.7) reacted.

1 3.9u: H-NMR (400 MHz, CDCl3) δ 5.62 – 5.21 (4H), 4.20 (dd, J = 8.0, 6.2 Hz, 1H),

4.08 – 3.97 (m, 1H), 3.63 (s, 3H), 3.30 – 3.25 (6H), 2.85 – 2.75 (m, 1H), 2.56 – 2.46 (m,

1H), 2.37 – 2.17 (m, 1H), 2.12 – 1.96 (4H), 1.50 – 1.32 (m, 2H), 1.32 – 1.15 (6H), 0.87 –

0.83 (12H), 0.06 – -0.05 (6H).

2 H-NMR (92 MHz, CHCl3) δ 2.25 (s, 2D), 1.99 (s, 2D), 1.61 (s, 2D).

+ + HRMS (EI) m/z 550.44045 (M·NH4 ) calcd for C29H52D6NO6Si found 550.44040.

1 3.9l: H-NMR (400 MHz, CDCl3) δ 5.58 – 5.19 (4H), 4.13 – 4.00 (2H), 3.69 – 3.58 (3H),

3.33 – 3.15 (6H), 2.77 – 2.58 (2H), 2.17 – 1.92 (5H), 1.52 - 1.34 (m, 2H), 1.32 – 1.15

(6H), 0.90 – 0.76 (12H), 0.06 – -0.06 (6H).

2 H-NMR (92 MHz, CHCl3) δ 2.23 (s, 2D), 1.94 (s, 2D), 1.58 (s, 2D).

+ + HRMS (EI) m/z 550.44045 (M·NH4 ) calcd for C29H52D6NO6Si found 550.44046.

(5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9-(dimethoxymethyl)-12-hydroxyheptadeca-

5,10-dienoate (3.10)

A solution of 3.9 (334.5 mg, 0.63 mmol) in THF (30 mL) was cooled to 0 oC and tetra-n-butylammoniumfloride solution (TBAF, 1.0 M in THF with 5% water, 10 mmol) was added. After stirring at 0 oC for 15 min, the cold bath was removed and the clear yellow solution was stirred at room temperature overnight under argon, during which it became clear brown. The reaction mixture was cooled to 0 oC and quenched by water (30 mL). The mixture was extracted with ethyl acetate. The combined organic layer was washed with water, then brine, and dried over anhydrous magnesium sulfate, filtered through a short silica gel column and concentrated by rotary evaporation to afford a slight turbid yellow oil. TLC analysis of the crude product showed that there were unreacted starting material 3.9 (15% ethyl acetate in hexanes, Rf = 0.3) and two product spots (50% ethyl acetate in hexanes, Rf = 0.35, 0.17). The crude product was purified by flash chromatography using 25%, 50% and 75% ethyl acetate in hexanes and the two product spots, designated as 3.10u and 3.10l corresponding to the spots Rf = 0.35, 0.17 respectively, were collected separately (overall yield 90% for3.10u and 3.10l; calculated based on the amount of starting material 3.9 reacted).

1 3.10u: H-NMR (400 MHz, CDCl3) δ 5.74 – 5.18 (4H), 4.28 – 4.02 (2H), 3.67 – 3.63

(3H), 3.39 – 3.19 (6H), 2.90 – 2.48 (2H), 2.33 – 2.16 (m, 1H), 2.16 – 2.05 (5H), 1.56 –

1.42 (m, 2H), 1.42 – 1.20 (6H), 0.90 – 0.83 (3H).

2 H-NMR (92 MHz, CHCl3) δ 2.25 (s, 2D), 1.99 (s, 2D), 1.59 (s, 2D).

+ + HRMS (EI) m/z 436.35398 (M·NH4 ) calcd for C23H38D6NO6 found 436.35393.

1 3.10l: H-NMR (400 MHz, CDCl3) δ 5.71 – 5.18 (4H), 4.29 – 4.02 (2H), 3.66 – 3.63

(3H), 3.39 – 3.18 (6H), 2.90 – 2.49 (2H), 2.32 - 2.04 (5H), 1.59 – 1.40 (m, 2H), 1.40 –

1.19 (6H), 0.91 – 0.78 (3H).

2 H-NMR (92 MHz, CHCl3) δ 2.24 (s, 2D), 1.95 (s, 2D), 1.58 (s, 2D).

+ + HRMS (EI) m/z 436.35398 (M·NH4 ) calcd for C23H38D6NO6 found 436.35394.

(5Z,10E)-2,2,3,3,4,4-d6-8-Acetyl-9-(dimethoxymethyl)-12-hydroxyheptadeca-5,10-

dienoic acid (isoLGE2-d6 acetal, 3.11)

To a solution of 3.10 (59.0 mg, 0.14 mmol) in water/methanol/THF (10 mL, 2/5/3, v/v/v) was added a solution of sodium hydroxide (16 mg/mL, 10 mL) in water/methanol/THF (2/5/3, v/v/v) at room temperature. The clear light yellow mixture was then magnetically stirred for 3 h, when TLC analysis (10% methanol in methylene chloride) confirmed the disappearance of starting material (3.10) and appearance of product 3.11 (Rf = 0.50). The reaction mixture was carefully acidified to pH 4 by addition of 2 M HCl, and the color of the solution changed from light yellow to colorless.

The mixture was extracted with ethyl acetate, and the combined organic layers were dried over anhydrous magnesium sulfate, filtered through a celite pad and concentrated by

rotary evaporation. The residue was purified by flash chromatography using 10%

methanol in methylene chloride to give 3.11 (51.6 mg, 0.13 mmol, 93%).

1 H-NMR (400 MHz, CDCl3) δ 5.73 – 5.17 (4H), 4.29 – 3.99 (2H), 3.36 – 3.18 (6H),

2.87– 2.50 (2H), 2.34– 1.90 (5H), 1.64 – 1.41 (m, 2H), 1.41 – 1.14 (6H), 0.85 (t, J = 6.3

Hz, 3H).

2 H-NMR (92 MHz, CHCl3) δ 2.25 (s, 2D), 1.98 (s, 2D), 1.60 (s, 2D).

+ + HRMS (EI) m/z 422.33832 (M·NH4 ) calcd for C22H36D6NO6 found 422.33836;

+ - - 403.29722 ([M-H ] ) calcd for C22H31D6O6 found 403.29710.

(5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9-formyl-12-hydroxyheptadeca-5,10-dienoic acid

(isoLGE2-d6, 3.12)

A solution of 3.11 (21.0 mg, 0.052 mmol) in acetic acid / water (3/1, v/v, 10 mL)

was magnetically stirred and the process of the reaction was monitored by TLC. After 3

h TLC analysis (10% methanol in methylene chloride) showed the product 3.12 (Rf =

0.52) and disappearance of the acetal 3.11 (Rf = 0.63). The solution was concentrated by rotary evaporation as well as high vacuum. The purity of final product 3.12 was estimated

by the ratio of integration of the aldehyde hydrogen (δ 9.42, 1H) to the three hydrogens (δ

0.82, t, 3H) of the terminal methyl group of the lower side chain.

+ + HRMS (EI) m/z 341.25935 ([M-H2O]·H ) calcd for C20H25D6O4 found 341.25941;

+ + 358.28590 ([M-H2O]·NH4 ) calcd for C20H28D6NO4 found 358.28599; 339.24479 ([M-

+ - - H2O] –H ) calcd for C20H23D6O4 found 339.24466.

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Appendix

1 Fig. A1 H-NMR spectrum of tert-butyl(1,1,2,2,3,3,4,4-d8-4-iodobutoxy)dimethylsilane (2.1)

2 Fig. A2 H-NMR spectrum of tert-butyl(1,1,2,2,3,3,4,4-d8-4-iodobutoxy)dimethylsilane (2.1)

13 Fig. A3 C-NMR spectrum of tert-butyl(1,1,2,2,3,3,4,4-d8-4-iodobutoxy)dimethylsilane (2.1)

1 Fig. A4 H-NMR spectrum of diethyl(4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)-2- oxoheptan-3-yl)phosphonate (2.2)

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm) Fig. A5 2H-NMR spectrum of diethyl(4,4,5,5,6,6,7,7-d -7-((tert-butyldimethylsilyl)oxy)-2- 8 oxoheptan-3-yl)phosphonate (2.2)

13 Fig. A6 C-NMR spectrum of diethyl(4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)-2- oxoheptan-3-yl)phosphonate (2.2)

1 Fig. A7 H-NMR spectrum ofdiethyl(4,4,5,5,6,6,7,7-d8-7-hydroxy-2-oxoheptan-3-yl)phosphonate (2.3)

2 Fig. A8 H-NMR spectrum ofdiethyl(4,4,5,5,6,6,7,7-d8-7-hydroxy-2-oxoheptan-3-yl)phosphonate (2.3)

61.6 61.2 31.5 31.0 25.5 25.0 24.5 24.0 23.5 f1 (ppm) f1 (ppm) f1 (ppm)

220 210 200 190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 -1 f1 (ppm) Fig. A9 13C-NMR spectrum ofdiethyl(4,4,5,5,6,6,7,7-d -7-hydroxy-2-oxoheptan-3-yl)phosphonate (2.3) 8

1 Fig. A10 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-(diethoxyphosphono)-6-oxoheptanoate (2.4)

Fig. A11 2H-NMR spectrum of methyl 2,2,3,3,4,4-d -5-(diethoxyphosphono)-6-oxoheptanoate (2.4) 6

13 Fig. A12 C-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-(diethoxyphosphono)-6-oxoheptanoate (2.4)

O D2 D2 C C OCH3 C C D2 H3CO O

H CO 3 2.5

1 Fig. A13 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-acetyl-7,7-dimethoxyhept-5-enoate(2.5)

O D2 D2 C C OCH3 C C D2 H3CO O

H CO 3 2.5

100

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm) 2 Fig. A14 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-acetyl-7,7-dimethoxyhept-5-enoate(2.5)

100

O D2 D2 C C OCH3 C C D2 H3CO O

H CO 3 2.5

101

33.0 25.0 24.5 24.0 23.5 23.0 f1 (ppm) f1 (ppm)

220 210 200 190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 -1 f1 (ppm) 13 Fig. A15 C-NMR spectrum of methyl 2,2,3,3,4,4-d6-5-acetyl-7,7-dimethoxyhept-5-enoate(2.5)

101

102

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

1 Fig. A16 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-9-(tert-butyldimethylsilyloxy)-6- (dimethoxymethyl)heptadeca-7,11-dienoate (2.8)

102

103

Fig. A17 2H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d -5-acetyl-9-(tert-butyldimethylsilyloxy)-6- 6 (dimethoxymethyl)heptadeca-7,11-dienoate (2.8)

103

104

1 Fig. A18 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9- hydroxyheptadeca-7,11-dienoate (2.9u)

104

105

2 Fig. A19 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9- hydroxyheptadeca-7,11-dienoate (2.9u)

105

106

1 Fig. A20 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9-

hydroxyheptadeca-7,11-dienoate (2.9l)

106

107

2 Fig. A21 H-NMR spectrum of (7E,11Z)-methyl 2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9-

hydroxyheptadeca-7,11-dienoate (2.9l)

107

108

1 Fig. A22 H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9- hydroxyheptadeca-7,11-dienoic acid (2.10)

108

109

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm) 2 Fig. A23 H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6-(dimethoxymethyl)-9- hydroxyheptadeca-7,11-dienoic acid (2.10)

109

110

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

Fig. A24 1H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d -5-acetyl-6-formyl-9-hydroxyheptadeca-7,11-dienoic acid (2.11) 6

110

111

2 Fig. A25 H-NMR spectrum of (7E,11Z)-2,2,3,3,4,4-d6-5-acetyl-6-formyl-9-hydroxyheptadeca-7,11-dienoic acid (2.11)

111

112

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

1 Fig. A26 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)hept-2-yn-1-ol (3.1)

112

113

2 Fig. A27 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)hept-2-yn-1-ol (3.1)

113

114

13 Fig. A28 C-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)hept-2-yn-1-ol (3.1)

114

115

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

1 Fig. A29 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)hept-2-yn-1-yl acetate (3.2)

115

116

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

2 Fig. A30 H-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)hept-2-yn-1-yl acetate (3.2)

116

117

61.5 18.0 17.5 24.0 23.5 31.0 30.5 30 f1 (ppm) f1 (ppm) f1 (ppm) f1 (ppm)

220 210 200 190 180 170 160 150 140 130 120 110 100 90 80 70 60 50 40 30 20 10 0 -1 f1 (ppm)

13 Fig. A31 C-NMR spectrum of 4,4,5,5,6,6,7,7-d8-7-((tert-butyldimethylsilyl)oxy)hept-2-yn-1-yl acetate (3.2)

117

118

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

1 Fig. A32 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-ynoate (3.3)

118

119

2 Fig. A33 H-NMR spectrum of methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-ynoate (3.3)

119

120

13 Fig. A34 C-NMR spectrum of methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-ynoate (3.3)

120

121

1 Fig. A35 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-enoate (3.4)

121

122

2 Fig. A36 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-enoate (3.4)

122

123

13 Fig. A37 C-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-hydroxyhept-5-enoate (3.4)

123

124

1 Fig. A38 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-bromohept-5-enoate (3.5)

124

125

2 Fig. A39 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-7-bromohept-5-enoate (3.5)

125

126

1 Fig. A40 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-8-(diethoxyphosphoryl)-9-oxodec-5-enoate (3.6)

126

127

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

2 Fig. A41 H-NMR spectrum of (Z)-methyl 2,2,3,3,4,4-d6-8-(diethoxyphosphoryl)-9-oxodec-5-enoate (3.6)

127

O D2 D2 C C C COOCH3 D2 H3CO

OCH3 3.7

128

1 Fig. A42 H-NMR spectrum of (5Z)-methyl 2,2,3,3,4,4-d6-8-acetyl-10,10-dimethoxydeca-5,8-dienoate (3.7)

128

O D2 D2 C C C COOCH3 D2 H3CO

OCH 3.7 3

129

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

2 Fig. A43 H-NMR spectrum of (5Z)-methyl 2,2,3,3,4,4-d6-8-acetyl-10,10-dimethoxydeca-5,8-dienoate (3.7)

129

130

1 Fig. A44 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12-((tert-butyldimethylsilyl)oxy)-9-

(dimethoxymethyl)heptadeca-5,10-dienoate (3.9u)

130

131

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

2 Fig. A45 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12-((tert-butyldimethylsilyl)oxy)-9-

(dimethoxymethyl)heptadeca-5,10-dienoate (3.9u)

131

132

1 Fig. A46 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-12-((tert-butyldimethylsilyl)oxy)-9- (dimethoxymethyl)heptadeca-5,10-dienoate (3.9l)

132

133

Fig. A47 2H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d -8-acetyl-12-((tert-butyldimethylsilyl)oxy)-9- 6 (dimethoxymethyl)heptadeca-5,10-dienoate (3.9l)

133

O D2 D2 C C C COOCH3 D2 H3CO

OCH3 OH 3.10

134

1 Fig. A48 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9-(dimethoxymethyl)-12-

hydroxyheptadeca-5,10-dienoate (3.10u)

134

O D2 D2 C C C COOCH3 D2 H3CO

OCH3 OH 3.10

135

2 Fig. A49 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9-(dimethoxymethyl)-12-

hydroxyheptadeca-5,10-dienoate (3.10u)

135

O D2 D2 C C C COOCH3 D2 H3CO

OCH OH 3 3.10

136

1 Fig. A50 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9-(dimethoxymethyl)-12-

hydroxyheptadeca-5,10-dienoate (3.10l)

136

O D2 D2 C C C COOCH3 D2 H3CO

OCH3 OH 3.10

137

1 Fig. A51 H-NMR spectrum of (5Z,10E)-methyl 2,2,3,3,4,4-d6-8-acetyl-9-(dimethoxymethyl)-12- hydroxyheptadeca-5,10-dienoate (3.10l)

137

138

0.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 -0 f1 (ppm)

1 Fig. A52 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9-(dimethoxymethyl)-12-

hydroxyheptadeca-5,10-dienoic acid (isoLGE2-d6 acetal, 3.11)

138

139

1 Fig. A53 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9-(dimethoxymethyl)-12-

hydroxyheptadeca-5,10-dienoic acid (isoLGE2-d6 acetal, 3.11)

139

O D2 D2 C C C COOH D2

O OH

isoLGE2-d6 (3.12)

140

1 Fig. A54 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9-formyl-12-hydroxyheptadeca-5,10-

dienoic acid (isoLGE2-d6, 3.12)

140

O D2 D2 C C C COOH D2

O OH

isoLGE2-d6 (3.12)

141

2 Fig. A55 H-NMR spectrum of (5Z,10E)-2,2,3,3,4,4-d6-8-acetyl-9-formyl-12-hydroxyheptadeca-5,10-

dienoic acid (isoLGE2-d6, 3.12)

141

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