DEVELOPMENT OF NOVEL SYNTHETIC ROUTES TO THE
EPOXYKETOOCTADECANOIC ACIDS (EKODES) AND THEIR
BIOLOGICAL EVALUATION AS ACTIVATORS OF THE PPAR FAMILY
OF NUCLEAR RECEPTORS
By
ROOZBEH ESKANDARI
Submitted in partial fulfillment of the requirements for
The Degree of Doctor of Philosophy
Thesis Advisor: Gregory P. Tochtrop, Ph.D.
Department of Chemistry
CASE WESTERN RESERVE UNIVERSITY
January, 2016 CASE WESTERN RESERVE UNIVERSITY
SCHOOL OF GRADUATE STUDIES
We hereby approve the thesis/dissertation of
ROOZBEH ESKANDARI
Candidate for the Ph.D degree *.
(signed) Anthony J. Pearson, PhD (Chair of the committee)
Gregory P. Tochtrop, PhD (Advisor)
Michael G. Zagorski, PhD
Blanton S. Tolbert, PhD
Witold K. Surewicz, PhD (Department of Physiology and Biophysics)
(date) 14th July, 2015
*We also certify that written approval has been obtained for any proprietary material contained therein.
I dedicate this work to my sister Table of Contents
Table of Contents ...... i
List of Tables ...... vi
List of Figures ...... vii
List of Schemes ...... ix
Acknowledgements ...... xi
List or Abbreviations ...... xiii
Abstract ...... xxiii
CHAPTER 1 General Introduction ...... 1
1.1 Discovery of prostaglandins ...... 1
1.2 The role of synthesis in the advancement of prostaglandins research ...... 3
1.3 Discovery of leukotrienes ...... 6
1.4 The role of synthesis in the advancement of leukotrienes research ...... 6
1.5 Non-enzymatic oxidation of PUFAs ...... 8
1.6 The role of synthesis in the advancement of LPO products ...... 10
1.7 Formation of LPO products ...... 12
1.8 Oxidative stress (OS) ...... 16
1.8.1 Formation of reactive oxygen species/reactive nitrogen species (ROS/RNS)
...... 17
1.8.2 Reactive oxygen species (ROS) ...... 17
i
1.8.2.1 Singlet oxygen ...... 17
1.8.2.2 Superoxide radical ...... 18
1.8.2.3 Hydrogen peroxide ...... 19
1.8.2.4 Hydroxyl radical...... 19
1.8.3 Reactive nitrogen species (RNS) ...... 19
1.8.3.1 Nitric oxide (NO.) ...... 19
1.8.3.2 Peroxynitrite (ONOO-) ...... 19
1.8.4 Reactivity of ROS/RNS...... 20
1.9. Biological roles of oxidized lipids ...... 22
1.9.1 Oxidized lipid and G protein-coupled receptors (GPCRs) ...... 23
1.9.2 Oxidized lipid and peroxisome-proliferator-activated receptors
(PPARs) ...... 24
1.10 References ...... 26
CHAPTER 2 Development of novel synthetic routes to the epoxyketooctadecenoic acids (EKODEs) ...... 31
2.1 Introduction ...... 31
2.2 EKODE structure and nomenclature ...... 34
2.3 EKODE-(E)-I family ...... 35
2.3.1 Retrosynthetic analysis of trans-EKODE-(E)- I ...... 36
2.3.2 Synthesis of trans-EKODE-(E)- Ia ...... 37
ii
2.3.3 Synthesis of trans-EKODE-(E)- Ib ...... 38
2.3.4 Retrosynthetic analysis of cis-EKODE-(E)- Ib ...... 40
2.3.5 Synthesis of cis-EKODE-(E)- Ib ...... 40
2.4 EKODE-(E)-II family ...... 42
2.4.1 Retrosynthetic analysis of trans-EKODE-(E)-II ...... 44
2.4.2 Synthesis of bifunctional conjunctive ylide ...... 45
2.4.3 Synthesis of trans-EKODE-(E)-II ...... 47
2.4.4 Divergent synthesis for diverse library of trans-EKODE-(E)-II
...... 49
2.4.4.1 Synthesis of alkyne-end aldehyde ...... 49
2.4.4.2 Synthesis of deuterated labeled aldehyde ...... 49
2.5 Experimental part ...... 52
2.5.1 General experimental details ...... 52
2.5.2 Syntheses ...... 54
2.6 References ...... 88
Chapter 3. Biological evaluation of EKODEs as potential activators of the PPAR family of nuclear receptors ...... 91
3.1 Peroxisome proliferator-activated receptors (PPARs) structure and activity ..... 91
3.2. The physiological functions of the PPARs ...... 95
3.2.1 Functional role of PPARα ...... 95
iii
3.2.1.1 Exogenous ligands (synthetic Xenobiotics) of PPARα ...... 95
3.2.1.2 Endogenous ligands (biological molecules) of PPARα ...... 96
3.2.2 Functional role of PPARɣ ...... 97
3.2.2.1 Exogenous ligands of PPARɣ ...... 98
3.2.2.2 Endogenous ligands of PPARɣ ...... 99
3.2.3. Functional role of PPARβ/δ ...... 101
3.2.3.1 Exogenous ligands of PPARβ/δ ...... 102
3.2.3.2 Endogenous ligands of PPARβ/δ ...... 102
3.3 EKODEs as potential endogenous metabolites ...... 103
3.4 Target reporter assay ...... 105
3.4.1 Bioluminescence ...... 105
3.4.2 Cell-based reporter assay ...... 105
3.4.3 Single luciferase reporter assay ...... 106
3.4.4 Luciferase enzyme mechanism ...... 107
3.4.5 Colorimetric β-galactosidase assay ...... 108
3.5 Experimental ...... 110
3.5.1 Materials ...... 110
3.5.2 Cell transient transfection assay ...... 110
3.5.3 Methods ...... 111
3.5.3.1 Seeding of COS-7 cells ...... 111
3.5.3.2 Transfection of COS-7 cells ...... 111
iv
3.5.2.3 Treatment of transfected cells ...... 112
3.5.2.4 Cell lysis ...... 112
3.5.2.5 Luciferase assay, data normalization ...... 112
3.6 Results and discussion ...... 113
3.7 Conclusion ...... 116
3.8 References ...... 118
Chapter 4. Thesis summary and future directions ...... 122
4.1 Thesis summary ...... 122
4.2 Future directions ...... 125
4.3 References ...... 127
APPENDIX NMR spectra of synthesized molecules ...... 128
BIBLIOGRAPHY ...... 181
v
List of Tables
Table 1.1 Physicochemical properties of reactive species (ROS/RNS), NA: data is not available ...... 20
Table 2.1 Reaction of sulfonium ylide and aldehydes, * yields were calculated based of amount obtained after purification ...... 50
Table 2.2 Reaction of phosphonium ylide and aldehydes.* yields were calculated based of amount obtained after purification ...... 51
vi
List of Figures
Figure 1.1 Non-enzymatic oxidized metabolites of PUFAs ...... 9
Figure 1.2 The molecular structure of linoleate and arachidonate esters ...... 13
Figure 1.3 Generation of different ROS/RNS and their subsequent reactions ...... 17
.- Figure 1.4 Standard electrode potential for superoxide anion radical (O2 ) and
. hydroperoxyl radical (HO2 ) ...... 21
Figure 1.5 Nernst equations can be used as an estimate to measure redox potential at known concentration and pH ...... 21
Figure 1.6 Lipid peroxidation products and their signaling through GPCR ...... 24
Figure 1.7 Lipid peroxidation products interact with PPAR and activate their gene expression machinery ...... 25
Figure 2.1 Linoleic acid oxidation through non-enzymatic reactions ...... 33
Figure 2.2 EKODE I and II structures and nomenclature ...... 34
Figure 3.1 PPARs motifs, conserved region in yellow, variable regions in red ...... 92
Figure 3.2 PPAR binding to ligand along with RXR binding to 9-cis retinoic acid (9cRA) forms a complex which provides an on- off-switch for gene expression ...... 93
Figure 3.3 Three main mechanisms proposed for negative regulation of transcription factors by PPARs ...... 94
Figure 3.4 Synthetic molecules and xenobiotics acting as PPAR-α ligands ...... 96
Figure 3.5 Endogenous (biological molecules) acting as PPAR-α ligands ...... 97
Figure 3.6 Synthetic molecules acting as PPARγ ligands ...... 99
vii
Figure 3.7 Endogenous molecules acting as PPARγ ligands ...... 100
Figure 3.8 Phospholipids which act as endogenous molecules acting as PPARγ ligands
...... 101
Figure 3.9 Synthetic molecules acting as PPARβ/δ ligands ...... 102
Figure 3.10 Endogenous molecules acting as PPARβ/δ ligands ...... 103
Figure 3.11 Illustration of the basic principle of the single luciferase reporter assay for
PPAR receptor ...... 106
Figure 3.12 Reaction catalyzed by firefly luciferase emits light through bioluminescence process ...... 108
Figure 3.13 Colorimetric assay of β-galactosidase using ONPG as the substrate ...... 109
Figure 3.14 Relative light units (RLU) determines activity of PUFAs and lipid metabolites on mouse PPARα by luciferase assay. 1X(5μM), 2X(10μM), 4X(20μM).
Wy14635 is the synthetic ligand ...... 113
Figure 3.15 Relative light units (RLU) determines activity of PUFAs and lipid metabolites on mouse PPARγ by luciferase assay. 1X(5μM), 2X(10μM), 4X(20μM) .
Troglitazone is the synthetic ligand ...... 114
Figure 3.16 Relative light units (RLU) determines activity of PUFAs and lipid metabolites on mouse PPARβ/δ by luciferase assay. 1X(5μM), 2X(10μM), 4X(20μM).
GW0742 is the synthetic ligand ...... 115
Figure 3.17 Relative light units (RLU) determines trans-EKODE-(E)-IIb activation of mouse PPARβ/δ in compare to GW0742 ...... 116
viii
List of Schemes
Scheme 1.1 General synthetic route for Corey lactone aldehyde ...... 4
Scheme 1.2 Synthetic route for prostaglandin E2and F2α ...... 5
Scheme 1.3 Synthetic route for leukotriene A4 (LTA4) ...... 7
Scheme 1.4 Synthetic route for 15-F2c-IsoP ...... 10
Scheme 1.5 Synthetic route for 8-epi-SC-Δ13-9-IsoF ...... 12
Scheme 1.6 Non-enzymatic formation of 15-F2c-IsoP ...... 14
Scheme 1.7 Non-enzymatic formation of 8-epi-SC-Δ13-9-IsoF ...... 15
Scheme 2.1 Non-enzymatic formation of EKODE-I family ...... 35
Scheme 2.2 Retrosynthesis analysis of trans-EKODE-(E)-I ...... 36
Scheme 2.3 Synthetic route for trans-EKODE-(E)-Ia ...... 38
Scheme 2.4 Synthetic route for trans-EKODE-(E)-Ib ...... 39
Scheme 2.5 Retrosynthesis analysis of cis-epoxy aldehyde ...... 40
Scheme 2.6 Synthetic route for cis-EKODE-(E)-Ib ...... 41
Scheme 2.7 Non-enzymatic formation of EKODE-II family ...... 42
Scheme 2.8 Different synthetic strategies in order to make intermediate X ...... 43
Scheme 2.9 Retrosynthesis analysis of trans-EKODE-(E)-II ...... 45
Scheme 2.10 Synthetic rout for bifunctional conjunctive ylide ...... 46
Scheme 2.11 General strategy for synthesis of trans-EKODE-II family ...... 47
Scheme 2.12 Synthetic route for trans-EKODE-(E)-IIa ...... 48
Scheme 2.13 Synthetic route for trans-EKODE-(E)-IIa an b ...... 48
ix
Scheme 2.14 Synthetic route for alkyne-end aldehyde ...... 49
Scheme 2.15 Synthetic route for deuterated-labeled aldehyde ...... 50
Scheme 4.1 At low temperature (0 oC), sulfonium ylide reacts faster than phosphonium
(k1>>k2) ...... 123
Scheme 4.2 Sulfonium ylide reaction is faster than phosphonium one ...... 124
Scheme 4.3 General strategy for phospholipid synthesis ...... 126
x
Acknowledgements
I would like to express my genuine appreciation to my advisor, Prof. Gregory P.
Tochtrop, for his inherent enthusiasm towards science education and sharing with me much of this feeling with me, and also encouraging me to take risks and pursue critical thinking as I grew into an independent scientist in his lab. All of these would not have been achieved without his patience.
I would like to thank the Department of Chemistry, Case Western Reserve
University and the National Science Foundation for financial support.
I also would like to thank my committee member, Prof. Anthony J. Pearson, Prof.
Michael G. Zagorski, Prof. Blanton S. Tolbert and Prof. Witold K. Surewicz, for precious time and effort put into my thesis.
I am so fortunate to study in such a great lab with individuals whose interest in revealing the fundamental aspects of science was inspiring. My best memories in the lab would not happened without: Dr. Emily C. Barker, Dr. Tonibelle Gatbonton-Schwager,
Dr. Yong Han, Jeremy P. Hess , Dr. Vasily A. Ignatenko, Dr. Qingjiang Li, Anna
Owensby, Jeffrey Rabinowitz, Dr. Sushabhan Sadhukhan, Chuan Shi, Elizabeth A.
Stewart, Dr. Brian S. Werry and Dr. Jianye Zhang. Especially, I am grateful to my lifelong friend Mohsen Badiee who happened to work in this lab for most of these years.
I am also thankful to Prof. Noa Noy and her students Dr. Liraz Levi and Mary K. Doud for their insightful comments and the lab space and instruments they provided for me in the Cleveland Clinic Lerner Research Institute. I am also grateful to Prof. Krzysztof
xi
Palczewski and his student Dr. Yuanyuan Chen for their great help when using the instruments in the Pharmacology Department at Case Western Reserve University.
Last but not least, my parents, Masoomeh and Mohammad Hassan, receive the deepest gratitude for teaching me how to enjoy being myself and providing me with love and support through all these years. I also express my sincere gratitude to my dear sister,
Ferdows, and brother-in-law, Jamshid, for being such great friends and inspiring me to do hard work.
xii
List of Symbols and Abbreviations
°C The degree Celsius
AA Arachidonic acid (arachidonate)
AF1 Activation function 1
AF2 Activation function 2
ANT Adenine nucleotide translocator
ATCC American Type Culture Collection
ATP Adenosine triphosphate
Azelaoyl PAF 1-O-hexadecyl-2-O-(9-carboxyoctanoyl)-sn-glyceryl-3- phosphocholine
α Alpha bp Boiling point
β Beta
βgal Beta-galactosidase
CBP CREB binding protein
CCD charge-coupled device
CDCl3 Deuterated chloroform
CH2Cl2 Dichloromethane
CH3CN Acetonitrile
10e12zCLA 10(E),12(Z)-Conjugated linoleic
xiii
COUP-TFII chicken ovalbumin upstream promoter-transcription factor
II
CREB cAMP response element binding
CVD Cardiovascular diseases d Doublet
D2O Deuterium oxide
DBD DNA binding domain
DCM Dichloromethane dd Doublet of doublets ddd Doublet of doublets of doublets
DEHA di-(2-ethylhexyl) adipate
DEHP di-(2-ethylhexyl)-phthalate
DHA Docosahexaenoic acid (docosahexaenoate)
DHKODE Dihydroxyletooctadecanoic acid
DMEM Dubelco's Modified Eagle Medium
DMSO Dimethyl sulfoxide
DNA Deoxyribonucleic Acid
DODE 9-12,-dioxo-10(E)-dodecenoic acid dt Doublet of triplets
δ delta
E0 Standard electrode potential
xiv
E’ Electrode potential
EI Electron impact
EKODE Epoxyketooctadecenoic acids
EPA Eicosapentaenic acid (Eicosapentaenoate) eq Equivalent
EtOAc Ethyl acetate
ESI Electrospray ionization
FA Fatty acid
FABP Fatty acid binding protein
FAS Fatty acid synthase
Fs Femtosecond
FT-ICR Fourier transform - Ion cyclotron resonance g Gram
16:0/18:1-GPC 1-palmitoyl-2-oleoyl-sn-glycerol-3-phosphocholine
GPCR G-protein coupled receptor
GFP Green fluorescent proteins
γ Gamma h hour(s)
HDAC Histone deacetylase
8(S)-HETE 8(S)-Hydroxy-(5Z,9E,11Z,14Z)-eicosatetraenoic acid
15(S)-HETE 15(S)-Hydroxy-5Z,8Z,11Z,13E-eicosatetraenoic
xv
HKODE Hydroxyketooctadecanoic acid
HMPA Hexamethylphosphoramide
4-HNE 4-Hydroxynonenal
HNF4 Hepatocyte nuclear factor 4
HO. Hydroxyl radical
. HO2 Hydroperoxyl radical
HODA 9-hydroxy-12-oxo-10(E)-dodecenoic acid
9-HODE 9-hydroxy-10E,12Z-octadecadienoic acid
13-HODE 13-hydroxy-9Z,11E-octadecadienoic
HOMO Highest Occupied Molecular Orbital
HPLC High-pressure liquid chromatography
9HpODE 9-hydroperoxy-10E,12Z-octadecadienoic acid
13HpODE 13-hydroperoxy-9Z,11E-octadecadienoic acid
HRMS High-resolution mass spectrometry
Hsp Heat shock protein
HWE Horner–Wadsworth–Emmons (reaction)
IC50 Half maximal inhibitory concentration
IsoF Isofurane
IsoP Isoprostane
IsoTx Isothromboxane
IsoLG Isolevuglandin
xvi
Keap1 Kelch-like ECH-associated protein 1 kcal/mol Kilocalorie per mole
KODDE Ketooctadecadienoic acid
L Laevorotatory
LA Linoleic acid (linoleate)
LBD Ligand binding domain
LDA Lithium diisopropyl amide
Leukotriene B4 5S,12R-dihydroxy-6Z,8E,10E,14Z-eicosatetraenoic acid lit. Literature
LPA Lysophosphatidic acid
LPO Lipid peroxidation
LT Leukotrienes
LTQ Linear trap quadrupole
Luc Luciferase
M Molar (concentration) m/z Mass-to-charge ratio
MAPK Mitogen-activated protein kinase m-CPBA Meta-chloroperoxybenzoic acid
Me Methyl
MEHP Monoethylhexyl phthalate mg Milligram
xvii
MHz Megahertz min Minute(s) mL Milliliter mm Millimeter mmol Millimole
Ms Millisecond nm Nanometer n-BuLi n-butyllithium
NCoR Nuclear receptor corepressor
NeuroK Neuroketal
NeuroP Neuroprostane
NFAT Nuclear factor of activated T-cells
NFκB Nuclear factor-κB
NitroLA 10-Nitrolinoleate
NMR Nuclear magnetic resonance
NO. Nitrogen oxide
. NO2 Nitrogen dioxide
NOS NO synthases
Nrf2 NF-E2 p45-related factor 2
NSAID Nonsteroidal anti-inflammatory drug
xviii
.- O2 Superoxide anion radical
OAP Oxidative addition product
OCP Oxidative cleavage product
4-ONE 4-Oxo Nonenal
ONPG ortho-Nitrophenyl-β-galactoside
ONP ortho-nitrophenol
ONOO− Peroxynitrite
− ONOOCO2 Nitrosoperoxycarbonate
ONOOH Peroxynitrous acid
OS Oxidative Stress
OUEA 11-oxoudec-9(E)-enoic acid oxLDL Oxidized low-density lipoproteins
PCC Pyridinium Chlorochromate
Pd Palladium
PG Prostaglandin
PGA1 Prostaglandin A1
PGA2 Prostaglandin A2
PGD2 Prostaglandin D2
PGI2 Prostaglandin I2 pH Potential hydrogen
xix
PFOS Perfluorooctanesulfonic acid
PPh3 Triphenylphosphine
PMA Phosphomolybdic acid ppm Parts per million
PPAR Peroxisome proliferator-activated receptors
PPL porcine pancreas lipase
PPRE Peroxisome proliferator response elements
PTM post- translational modification
PUFA Polyunsaturated fatty acid
RGS4 Regulator of g-protein signaling 4
RLU Relative light units
RNA-seq RNA Sequencing
RNS Reactive nitrogen species
ROS Reactive oxygen species
RT-PCR Reverse transcription polymerase chain reaction
RXR Retinoid X receptor
SMRT Silencing mediator of retinoid and thyroid hormone receptor
SRC1 Steroid receptor co-activator 1
SRSA Slow reacting substance of anaphylaxis
xx
STAT Signal transducer and activator of transcription t1/2 Half-life t-EKODE-Ia trans-EKODE-( E)-Ia t-EKODE-Ib trans-EKODE-( E)-Ib t-EKODE-IIa trans-EKODE-( E)-IIa t-EKODE-IIb trans-EKODE-( E)-IIb
THF Tetrahydrofuran
Torr millimetre of mercury
TLC Thin-layer chromatography
TX Thromboxane
TP Thromboxane A2-Prostanoid
UV Ultraviolet
δ Delta, chemical shift
λmax Lambda maximum (wavelength)
μg Microgram
μm Micrometer (distance)
μs Microsecond
μM Micromolar (concentration)
WADA World Anti-Doping Agency
ω Omega
xxi
X-ray X-radiation
− X³Σg Triplet ground state
xxii
DEVELOPMENT OF NOVEL SYNTHETIC ROUTES TO THE
EPOXYKETOOCTADECANOIC ACIDS (EKODES) AND THEIR
BIOLOGICAL EVALUATION AS ACTIVATORS OF THE PPAR FAMILY
OF NUCLEAR RECEPTORS
Abstract
By
ROOZBEH ESKANDARI
Current drug screening processes help the scientific community develop therapeutics for a variety of specific targets and diseases, but the ability to identify an endogenous molecule or metabolite as a novel pharmacophore still remains an exciting area. In fact, identifying the putative target of endogenous metabolites can provide a deeper layer of understanding regarding cell physiology and regulatory feedback mechanisms. Furthermore, endogenous metabolites play a key role in developing diseases biomarkers and help to develop diagnostic tools.
Moreover, new scaffolds of metabolites can provide possibilities for developing novel therapeutics to treat related diseases, since their action is mostly unique toward a specific target in the cell.
In this regard, oxidized lipid metabolites formed by non-enzymatic reactions have not been systematically studied as signaling molecules. Among the variety of polyunsaturated fatty acids (PUFAs), a family of linoleic acid
xxiii metabolites called EKODEs has shown activities as endogenous metabolites.
Indeed, these molecules exhibit activities such as modulating corticosterone and aldosterone production levels. They also demonstrate antioxidant effects through the Nrf2-Keap1 signaling pathway.
As enthusiasm for understanding these physiological roles increased among different groups, the lack of synthetic methods for preparation of these molecules became more noticeable. This thesis represents a general strategy for providing a highly efficient synthesis for EKODE metabolites. Structurally, this family of metabolites posseses flanking α, β-unsaturated and epoxy functionalities centered on a ketone group. The synthetic method relies on developing a bifunctional intermediate containing two reactive ylides that can be used for generation of epoxy and olefin functionality in a sequential order. In fact, the two ylides kinetically differentiate between their reactivity with various electrophiles, such as aldehydes. This strategy can be used by others as a pipeline to generate a diverse family of active lipid metabolites with similar functionality.
This thesis also contains an initial biological evaluation of EKODEs, along with a number of other lipid peroxidation metabolites on the PPAR family of nuclear receptors. Our data suggest that these endogenous metabolites can regulate PPARβ/δ subtype at low micromolar concentration ranges. These receptors are specifically involved in the development of different chronic diseases, such as obesity, atherosclerosis and cancer. The information presented in
xxiv this thesis can be used as the stepping stone to reveal the chemical physiology of these endogenously formed metabolites.
xxv
Chapter 1: General Introduction
Attempts to understand human physiology date back to at least 420 BC, to the time of Hippocrates when he proposed the idea of the four basic substances or humors. In
19th century, the number of physiological studies started to grow more rapidly, and in the
1850s a French physiologist named Claude Bernard coined the word Milieu intérieur
(internal environment), which was expanded to homeostasis by an American physiologist
Walter B. Cannon in the 1920s. Cannon suggested that the human body is an open
system, exposed to dynamic changes from the environment, and it functions through
cooperating mechanisms to maintain a steady-state.1,2
Accordingly, scientists started to appreciate describing physiological phenomena
in a more detailed manner from a chemical point of view. In the 1930s, British
physiologist Henry H. Dale proposed the concept of autopharmacology while studying
acetylcholine interaction with muscles.3 Consequently, his discovery provided the foundations for his students to search for endogenous chemical substances that are involved in the regulation of physiology.
1.1 Discovery of prostaglandins
Among Dale’s students, Ulf von Euler, with his expertise and passion for science, discovered a lipid-soluble organic acid with hypotensive and muscle-stimulating actions in human semen. He called these molecules prostaglandin (PG), since the initial thought was that these molecules derived from prostate gland secretions. However, later it was found that the molecules secrete from seminal vesicles. Subsequently, similar molecules were discovered in other tissues with diverse physiological functions.4
1
The study in the field of PGs was partly halted by World War II. Eventually, in
1945 at a meeting at the Physiological Society of Karolinska Institute in Stockholm, von
Euler persuaded Sune Bergström, a lipid chemist, to analyze the structure of PGs. Due to
the lack of sensitive analytical methods, they required a large quantity of vesicular glands
from across the globe to obtain a sufficient amount of PGs to characterize the structures.
Bergström took advantage of stainless steel countercurrent extraction to purify the
samples, and he recognized them as unsaturated hydroxy acids in 1949.5
Improvements in chromatography techniques provided an ideal advantage for
6 purification and obtaining crystal forms of PGE1 and PGF1α. UV-spectroscopy and IR were among the first analytical methods used to determine the structure and quantity of these molecules. Subsequently, more sensitive techniques such as mass spectrometry and its combination with gas chromatography, developed by Ragnar Ryhage, provided the opportunity to characterize the formula and structure of PGs in pico- and nanogram
quantities in a variety of tissues.7
David van Dorp at Unilever research laboratory collaborated with Bergström to
investigate the transformation of isotopically labeled dihomogamma-linoleic acid to
PGE1 and PGF1α with homogenates of sheep glands. This result was critical, because it
confirmed that prostaglandins are oxidized forms of polyunsaturated fatty acids
(PUFAs).8 Subsequently, more detailed studies done with chemical and enzymatic
reactions provided ample information about the main scaffold, functional groups and
their arrangement in the structure of PGs. For example, oxidative ozonolysis mediated
fragmentation of the molecule to smaller pieces that could then be identified with other
2
9 methods, such as mass spectrometry. In addition, X-ray analysis of derivatives of PGF1α
by Sixten Abrahamsson revealed the first absolute configuration of PGs.10
The increasing number of biological applications of PGs, such as being local
hormones, inflammation and pain mediators, vasomotor regulators and neuromodulators
in animals, urged scientists to look for enriched sources. A study of natural products
accidentally discovered that Gorgonia coral contains about 1.0-1.5% of derivative of
11 PGA2. Consequently, the Upjohn Company used coral as the main source to obtain different forms of prostaglandins. Although natural sources provided a variety of PG
structures, in the late 1960s the remarkable work of Elias J. Corey established efficient
synthetic approaches to generate larger quantities of these molecules. In the next section,
the advances in the field of prostaglandins through synthesis are discussed.
1.2 The role of synthesis in the advancement of prostaglandins research
The major hurdles in the synthesis of PGs were that these molecules are quite unstable and they exist in a variety of scaffolds. One of the key discoveries in Corey’s work was the design of a single intermediate, commonly known as the Corey lactone aldehyde.12 In other words, this intermediate was not only used to synthesize PGs, it was
also applied to the generation of PG analogues that were the main focus of the
pharmaceutical industries. This general synthetic route for Corey lactone aldehyde is
3
shown in Scheme 1.1.
Cl CN MeO NaH, THF KOH, H2O/ DMSO Cl o MeOCH Cl OMe Cu(BF4)2, 0 C 2 CN THF, -55 oC
COOH MeO MeO o m-CPBA, NaHCO3 1. NaOH, H2O, 0 C KI3, NaHCO3 o O O 2. CO2 OMe H2O, 0 C O HO
O O O O O O o 1. Ac2O, Py 1. BBr3, CH2Cl2, 0 C I n o OMe 2. ( -Bu)3SnH OMe 2. CrO3, 2Py, CH2Cl2, 0 C O ALBN,Benzene HO AcO AcO
Scheme 1.1 General synthetic route for Corey lactone aldehyde
Next, the Corey lactone aldehyde afforded olefin bonds after going through several steps including Wittig and Horner-Wadworth-Emmons (HWE) coupling.13 In
Scheme 1.2, the procedures for PGE2 and PGF2α are presented. Similar strategies can be applied to other members of PG1’s, PG2’s and PG3’s. These syntheses facilitated the evaluation of the physicochemical properties of PGs and assessment of their biological implications on a worldwide scale.
4
O O O O O O P (MeO)2 n-C 5H11 Zn(BH4)2, DME
O NaH,DME n-C5H11 AcO AcO O Corey lactone aldehyde
O O O O 1. of Separation 1. Dibal-H, toluene, -60 oC diastereomers n-C5H11 - , 2. COO 2. K2CO3 MeOH n-C5H11 AcO Ph3P OH 3. DHP,TsOH THPO DMSO CH2Cl2 OTHP
O COOH
HO OH HO PGE2 COOH
THPO OTHP HO COOH
HO OH
PGF2α
Scheme 1.2 Synthetic route for prostaglandin E2 and F2α
Meanwhile, many novel syntheses and purification methodologies were developed in order to achieve a higher yield for the generation of PGs. For example, in the preparation of PGs’ stereocenters, molecular robots opened new areas in the total synthesis. These robot-like assemblers act in multi-step processes, interacting with
5
reactants and reagents to provide a stereoselective environment to obtain the enantiomerically pure products.14,15
1.3 Discovery of leukotrienes
The research on metabolites of PUFAs became one of the major foci at the
Karolinska Institute. In 1977, Beng I. Samuelsson discovered that during the
inflammatory response in leukocytes, arachidonic acid transforms to novel scaffolds
called leukotrienes (LT). More detailed studies by his postdoctoral researcher, Pierre
Borgeat, proved that bioconversion of arachidonic acid to hydroperoxide 5-HPETE is the
16 key step in formation of LTA4. Consequently, LTA4 converts to other forms of LTs: reaction with hydrolase enzyme forms LTB4, reaction with glutathione forms LTC4 and etc. These metabolites play a key role in hypersensitivity reactions, such as asthma and
inflammation.17
1.4 The role of synthesis in the advancement of leukotrienes research
In collaboration with Corey’s lab for synthesizing LTs in large quantities,
Samuelsson mainly focused on studying this newly found family of oxidized lipids. LTA4
was first synthesized as the racemate and soon after, its chiral form generated from D-(-)-
6
ribose.18 (Scheme 1.3)
1. Ph3P COOEt OAc OBz O BzO OH PhCOOH, DME, ∆ BzO Zn/Hg, HCl COOEt Et O 2. Ac O,H SO 2 BzO OBz 2 2 4 OBz
OAc OTs BzO 1. H2, Pd/C. MeOH BzO K2CO3, MeOH COOEt COOMe 2. HCl, MeOH OBz OBz 3. TsCl,Py
1. CrO3.Py,CH2Cl2 Li 2. OEt H H O O THF, -78 oC COOMe COOMe HO o O 3. MsCl, Et3N, CH2Cl2, -45 C H H o 4. pH 7, -45 C to 0 0C
o O 1. HMPA, -78 C COOMe 2. n-BuLi, THF, -78 oC
n-C5H11 Ph3P LTA 4
Scheme 1.3 Synthetic route for leukotriene A4 (LTA4)
The stereocontrolled synthetic strategies for LTs were used to confirm the
structure and the link between 5-HPETE and other metabolites such as LTA4 and LTB4.
In addition, the biogenic components of the slow reacting substance of anaphylaxis
(SRSA) were proved by synthesis of the peptidic LTC4 and etc.
With the aid of novel synthetic approaches, a myriad of oxidized forms of PUFAs
were characterized and studied. Meanwhile, isotope-labeling and advances in molecular biology provided plenty of information for understanding the origin of oxygen atoms in 7
the oxidized lipids.19 Accordingly, the synthetic intermediates were necessary to
unambiguously establish the absolute stereochemistry and study the mechanisms of enzymatic reactions involved in the biosynthesis of PGs, LTs and etc.
1.5 Non-enzymatic oxidation of PUFAs
The long-standing paradigm was that the enzymatic reactions were the only source of oxidized lipids production. In the 1960s, scientists observed examples of oxidized lipids derived from oxidation of PUFAs without the aid of enzymes. 20 Next, in
the late 1970s Robert G. Salomon discovered that as an enzymatic intermediate PGH2 can
rearrange spontaneously to generate different forms of PGs and thromboxanes (TXs).21 In
the 1990s L. Jackson Roberts and coworkers discovered a number of other oxidized
PUFAs called isoprostanes (IsoPs), neuroprostanes (NeuroPs), isothromboxanes (IsoTxs),
isolevuglandin (IsoLGs), neuroketals (NeuroKs) and isofuranes (IsoFs) that were produced through reactive oxygen species (ROS).22-27 (Figure 1.1)
In general, the formation of these oxidized metabolites, either spontaneously or
through ROS, follows the conventional rules of chemistry. These processes allow the
generation of greater numbers of scaffolds while forming racemic mixtures of
stereoisomers. The non-enzymatic reactions on lipids are generally called lipid
peroxidation (LPO).
8
HO HO OH COOH COOH
HO OH HO 7-F4t-NeuroP 15-F2t-IsoP O
COOH O O OHC OH OH
15-A2-IsoTx Iso[4]LGE4(12-E2-IsoK)
OHC COOH HO COOH O OH
O OH OH -NeuroK) IsoFs Iso[9]LGD4(17-D4
Figure 1.1 Non-enzymatic oxidized metabolites of PUFAs
The fact that these metabolites exist in significant amounts in vivo and exhibit
biological activities encouraged the science community to discover their cognate
receptors. Biological activity of LPO products is tissue-specific. Many of these activities are attributed to signaling through Thromboxane A2-Prostanoid (TP) receptor, which is a
G protein-coupled receptor (GPCR). Also, they function through the nuclear receptors,
such as peroxisome proliferator-activated receptors (PPARs).28 In addition to the
biological activities on a variety of lipid targets, LPO products such as F2-IsoPs, F4-
NeuroP and isofurans were used as the gold standard of oxidative stress.29-31
Indeed, the application of LPO products in biological systems triggered a strong
research effort to develop synthetic methods for generating these molecules. Many of the
9
syntheses are similar to the ones developed for products of enzymatic reactions with modified steps towards developing new stereochemistry.32
1.6 The role of synthesis in the advancement of LPO products
One of the well-studied classes of LPO products are IsoPs. The first total syntheses for this family mostly relied on biomimetic routes through radical cyclization reactions. Interestingly, these methods were developed in 1984, even before the discovery of these molecules in the process of developing synthesis for enzymatic reaction products.33 In the 1990s, multiple research groups developed a variety of strategies to generate all-cis-Corey lactone aldehyde for the synthesis of specific stereoisomers.34 One
35 of these strategies for synthesis of 15-F2c-IsoP is shown in Scheme 1.4.
COOMe
O COOMe 1. NaBH4 PhSeCH(COOMe)2 O 2. NaOH/MeOH, DMSO 140 oC Benzene, Sunlamp O 300 W O SePh 3. LiHMDS, PBzCl
COOMe O COOMe O 5 steps O H2O2, acetone-water, PPTS
PBzO SePh O (see Scheme 1.2) PBzO
HO COOH
HO OH
15-F2c-IsoP
Scheme 1.4 Synthetic route for 15-F2c-IsoP 10
At high oxygen concentration in tissues, arachidonic acid oxidation mainly switches to a different class of molecules known as isofurans (IsoF). In Scheme 1.5, synthesis for 8-epi-SC-Δ13-9-IsoF is presented. The key intermediate was a diol epoxide that was obtained through two different Sharpless asymmetric reactions. Next, specific arrangement of the tetrahydrofuran ring and hydroxyl groups was acquired through an epoxide cyclization cascade. In addition, the neat product of 5-exo-tet cyclization was used through Mitsunobu inversion to gain access to the other stereoisomers of isofurans.36
11
Br OH 1. NaI, CuI, K2CO3 Sharpless H asymmetric 1. Et3N, TMSCl OH epoxidation 2. OH 2.LiAlH4 H O O O O O OH
Br2
O 1. TBSCl O O 2. BuLi, BF OEt OH OSO2Ph 3. 2 OH O H HO 1. PhSO2Cl. EtN3 K2CO3 HO CN 2. AD-mix-α O O O O O O
HO HO NC 1. TBDPSCl - 1. P-2 Ni, H2 n O 2. C5H11MgBr + O TBSO 2. H3O HO CN O O CHO
TBDPSO NC HO HOOC
O 2 steps O TBDPSO HO
R2 R2 R 1 R1 R1 = OH, R2 = H
R1 = H, R2 = OH
Scheme 1.5 Synthetic route for 8-epi-SC-Δ13-9-IsoF
1.7 Formation of LPO products
12
PUFAs, as one of the major components of cell membranes, are subject to
modifications with reactive species. The reactions mostly occur through radical mechanisms, building a family of products distinct from enzymatic ones. The radical chain reactions have been categorized in three different major steps: (1) Initiation: the
key event which involves hydrogen-abstraction by radical or oxidant. (2) Propagation:
oxygen or nitrogen addition to carbon-centered radicals, followed by fragmentation and the rearrangement of atoms and bonds. (3) Termination: when radicals combine with each other and stop reacting by forming non-radical products.
7 COOR 11 10 C5H11 (CH2)7COOR 13 Linoleate,LA;18:2 Arachidonate,AA;20:4
COOR COOR
Docosahexanoate,DHA;22:6 Eicosapentaenoate,EPA;20:5
Figure1.2 The molecular structure of major oxidizabale PUFA esters.
In this regard, polyunsaturated fatty acids (PUFAs), such as linoleate (LA), arachidonate (AA), eicosapentaenoate (EPA), docosahexaenoate (DHA) and their esters are predisposed to undergo peroxidation. This is due to the C-H bonds at the bis-allylic positions having lower bond dissociation enthalpy. In other words, the bis-allylic C-H bonds are the weakest bonds in these molecules, and the hydrogen atoms at these positions are favorably abstracted during oxidation.37 Next, the peroxidation reaction in
13 generation for the two nonenzymatic reaction products 15-F2c-IsoP and 8-epi-SC-Δ -9-
IsoF is discussed.
13
O O ROS COOH COOH n n-C5H11 -C5H11 H H H H 3 3
radical attack site
O O O O O O O O COOH COOH n n-C H -C5H11 5 11 3 3 disrotatory (racemic) 11-HpETE cyclic peroxide process
COOH COOH HO O O O O O O O
H H
syn to cyclic peroxide
HO
steps COOH O COOH O HO OH OOH 15-F c-IsoP 15-G2c-IsoP 2 Scheme 1.6 Non-enzymatic formation of 15-F2c-IsoP
The postulated mechanism for formation of the lipid peroxidation product 15-F2c-
IsoP is summarized in Scheme 1.6. The first step involves the conversion of arachidonic acid (AA) to peroxyl form. Following this, the radical reacts internally with the neighboring double bond, producing the short-lived intermediate cyclic peroxide. Next, the two radical π-orbitals combine to form a new σ-bond, thus forming the cyclopentane ring. This process undergoes disrotatory ring formation based on woodward-hoffmann rules for pericyclic reaction. The inward rotation can happen at the syn or anti stereochemistry comparing to the cyclic peroxide. In this case the syn stereoisomer forms and eventually breaks down to F-ring IsoP through spontaneous reactions.38
14
Similarly, at higher level of oxygen 8-epi-SC-Δ13-9-IsoF is derived from arachidonic acid. Once the cyclic peroxide is formed, it undergoes a facile 1,3-SHi reaction to form
diepoxy hydroperoxide. The formation of diepoxide is competing with the formation of
the cyclopentane ring in IsoP.
O O ROS COOH COOH n n-C5H11 -C5H11 H H H H 3 3
radical attack site
O O O O COOH COOH n n-C H -C5H11 5 11 3 3 1,3-SHi (racemic) 11-HpETE
O O O O O O O O COOH H2O COOH n-C5H11 n-C5H11 3 3 diepoxy peroxide
HO HO O HO OH O O O HO OH COOH COOH n-C5H11 n-C5H11 3 3
HO
O COOH OH OH
8-epi-SC-∆13-9-IsoF Scheme 1.7 Non-enzymatic formation of 8-epi-SC-Δ13-9-IsoF
Next, oxygen adds to the molecule on the newly-generated radical to form diepoxy
hydroperoxide. After hydrolysis of the epoxide ring to form the regioisomeric expoxy
diols, an intramolecular nucleophilic ring opening of the epoxide through 3-exo
15
cyclization generates the tetrahydrofuran ring. (Scheme 1.7) The order of some steps may
be different from the mechanism shown, but the products will not change.39,40
Because LPO is one of the downstream reactions of oxidative stress, there has been an increasing focus on understanding the role of pathways related to ROS/RNS.41
1.8 Oxidative stress (OS)
Cells are capable of providing diverse mechanisms to maintain balance between the
production and consumption of the species of redox reactions. The discrepancy in these
regulatory systems can lead to the accumulation of more oxidants, which cause oxidative
stress. In other words, oxidative stress originates either from higher levels of oxidant formation or from the inhibition of antioxidant protective mechanisms.42
Together, oxidative stress and nitrosative stress have been identified as the major
contributors to the pathogenesis and pathophysiology of aging, cancer and chronic
disease.43 They primarily provide reactive species and radicals that are capable of
chemically modifying central components of the cell and producing molecules with
deviated functions. However, oxidative stress has also been recognized as a regulatory mechanism for homoeostasis. Indeed, cells take advantage of these mechanisms to perform necessary radical reactions degrade old machineries and regenerate new constituents. For example, the human immune system utilizes the excessive production of reactive oxygen and nitrogen species to destroy pathogens through a process termed the oxidative burst. 41,44
16
1.8.1 Formation of reactive oxygen species/reactive nitrogen species (ROS/RNS)
Oxidative stress is mainly caused by reactive oxygen species; similarly, nitrosative
stress is attributed to reactive nitrogen species. In physiological conditions, the reactive
species are formed during the mitochondrial electron transport or through different
enzymatic activities and free metals in the cells.44 Once the reactive species are
generated, they can react with each other to form a variety of combinations that can be
more reactive than the original ones. 45(Figure 1.3) As a signaling entity in the cell, their chemical physiology is dictated by different physiochemical factors such as concentration, half-life (t1/2), rate of diffusion across membranes and standard electrode
potential (E0).
1 O2 - ROO / RO RH ν e h RH - - e - e OH ROS O2 O2 H2O2 H+
Fe2+ Fe3+
+ - H NO ONOO ONOOH NO2 RNS CO 2
ONOOCO2
- CO3 Figure1.3 Generation of different ROS/RNS and their subsequent reactions
1.8.2 Reactive oxygen species (ROS)
1.8.2.1 Singlet oxygen
Atmospheric oxygen (O2) exists in a biradical form in which both electrons in the
− open-shell triplet ground state O2 (X³Σg ) have the same spin in two degenerate
17
antibonding πg-orbitals. This feature makes O2 “spin restricted” and not highly
chemically reactive. However, once energy is provided, the spin restriction is
overcome and the molecule is excited to other electronic states of antiparallel spins or singlet spins. In excited forms the spin can be in two different arrangements: O2 (a¹Δg)
+ (non-radical) and O2 (b¹Σg ) (more reactive free radical), the latter of which is less
46 stable and relaxes quickly to the lowest lying excited state O2 (a¹Δg). O2 (a¹Δg) is
considered the most biologically relevant reactive oxygen species. With a half-life of
47 1 μs, O2 (a¹Δg) has sufficient time to diffuse into various targets within the cell.
1.8.2.2 Superoxide radical
Singlet oxygen can lose its spin restriction in a facile process of one-electron
.- reduction to form the superoxide anion radical (O2 ), which is more reactive than
.- triplet oxygen. With a half-life less than 1 μs, O2 has minimal time for diffusion,
.- reaching a few micrometers from the site of generation. O2 acts a Brønsted base and
+ . reacts with H , forming hydroperoxyl radical HO2 . The acidic form is more stable
and hypothetically more permeable through membranes. The pKa for this acid-base
conjugate is around 4.88, which means that compartments with lower pH, such as the
mitochondrial intermembrane space, peroxisomes and lysosomes, would
.- accommodate the acidic molecular form. O2 is primarily a reducing agent. However,
48,49 .- the acidic form acts as an oxidant. Although O2 is not capable of modifying
macromolecules, it acts as a major source of oxidative stress. Specifically, it functions
.- . as a reducing species, which generates many other strong oxidants. O2 / HO2 can
produce hydrogen peroxide (H2O2), which can be decomposed through the Fenton reaction (presence of metal ions such as iron or copper) to form hydroxyl radical
18
. . - (HO ). In addition, superoxide anion radical reacts with NO and forms ONOO , an
extremely reactive RNS.
1.8.2.3 Hydrogen peroxide
Hydrogen peroxide (H2O2) is a weak acid without any unpaired electrons (a non- radical) that has a short half-life 10 μs because of the activities of detoxifying enzymes such as catalase and peroxidase. It has enhanced diffusion across long distances and membranes, although it is likely that this oxidant is less membrane permeate than a gas such as nitric oxide.
1.8.2.4 Hydroxyl radical
Hydroxyl radical (HO.) is the predominant source of oxidative stress that can
damage a wide variety of macromolecules in the cell. It possesses a half-life less than
1 fs. It is mainly biosynthesized through metal centers and is typically referred to as
Fenton-derived, named after Fenton who studied hydrogen peroxide production in
19th century.50
1.8.3 Reactive nitrogen species (RNS)
1.8.3.1 Nitric oxide (NO.)
Nitrogen oxide (NO.) is the product of the enzymes called NO synthases (NOS),
from the reaction of L-arginine and oxygen to produce citrulline and NO. It has a
half-life of 3-5s and is capable of traveling 50–200 μm from its production
source..51,52
1.8.3.2 Peroxynitrite (ONOO-)
Peroxynitrite forms through combining nitrogen oxide and superoxide anion
.- − radical (O2 ). ONOO is a strong nucleophile and exhibits a half-life of about 10 ms.
19
Its protonated form, peroxynitrous acid (ONOOH), is an extremely strong oxidant
(pKa= 6.8). It is capable of directly reacting with organic moieties or undergoing
. . homolysis to nitrogen dioxide (NO2 ) and a hydroxyl radical (HO ). In addition,
− ONOO can react with CO2 (1.3 mM in plasma) to form nitrosoperoxycarbonate
− − (ONOOCO2 ). ONOOCO2 eventually breaks down to two strong and short-lived
.- oxidant radicals, a carbonate radical (OCO2 ) and a nitrogen dioxide as a pair of
caged radicals. 53,54
1.8.4 Reactivity of ROS/RNS
A summary of physicochemical properties of major ROS/RNS is presented in
Table 1.1. These reactive species are formed endogenously through both enzymatic and spontaneous reactions and provide signaling roles in the body. 41
Species In vivo (t1/2)(s) reactivity reaction Standard electrode
molarity potential(v)
1 -6 - 0 O2 NA 10 oxidant 1∆ E =+0.64 gO2 + e O 2
.- -11 -10 -6 - 0 O2 10 -10 10 reductant e E =-0.35 O2 + O 2
-7 -5 -3 - 0 H2O2 10 10 -10 oxidant + E =+0.80, H2O2 + H + e HO + H2O
E0’=+0.39 (pH 7.0)
. -15 -9 + - 0 HO 10 10 oxidant HO + H + e H O E =+2.73, 2 E0’=+2.31 (pH 7.0)
NO. NA 3-5 oxidant + - E0=-0.11 NO + H + e HNO
ONOO- 10-9 10-2 oxidant E0’=+1.4 (pH 7.0) ONOO + 2H + e NO2 + H2O
. 0 NO2 NA NA oxidant - E =+1.04 NO + e NO 2 2
Table1.1 Physicochemical properties of reactive species (ROS/RNS), NA: data is not available.
20
Many of these reactive species are in equilibrium in water. As a result, pH has an
impact on their redox activity. In this regard, the standard electrode potential can provide
valuable information for ROS/RNS species and determine their role in redox biology,
either as oxidants or reductants.
.- For example, under physiological conditions, the superoxide anion radical (O2 ) tends
. to be a reducing agent in comparison to the hydroperoxyl radical (HO2 ). This may not be
readily concluded from the given standard electrode potential (E0) values.
- + - ' + 0 O + 2H + e H O E =+0.97(pH=7) HO2 + H + e H2O2 E =+1.46 2 2 2
- - + e 0 0 O2 + H + HO E =-0.03 O + e O2 E =-0.35 2 2
.- Figure 1.4 Standard electrode potential for superoxide anion radical (O2 ) and hydroperoxyl radical
. (HO2 )
In fact, the Nernst equation can provide a more accurate estimate in which both the
concentration of the reactive species and the pH are necessary components.55(Figures 1.4
& 1.5)
- Ox + mH+ + ne Red
Ox 10 -(m)(pH) E = E0 + 0.059/n log X Nernst equation Red
Figure 1.5 Nernst equations can be used as an estimate to measure redox potential at known concentration and pH
Although the Nernst equation can give an approximate idea about the inherent redox
activity of different species, it can hardly be used to investigate the level of reactivity.
Reactivity is more attributed to kinetics, meaning that the level of reactivity for these
species in the complex network of the cell is mostly dependent on kinetic parameters. 21
The half-life (t1/2) of the reaction for the species provides more valuable information than
thermodynamic properties such as the Nernst equation.56
1.9. Biological roles of oxidized lipids
Although originally scientists in the field were more interested in oxidized lipids derived from enzymatic reactions, non-enzymatic products also became important to the community in the 1980s. The change mainly occurred due to the many LPO products
detected in vivo in both clinical and experimental models. Indeed, the characterization of
adducts to these molecules has been linked to the onset and progression of several
pathological states associated with oxidative stress.43
The reported concentration for oxidized lipids in the cell is often in the nanomolar
range, raising an important question regarding the mechanism by which oxidized lipids
can exert their signaling roles. In fact, one of the crucial features contributing to their function is the existence of reactive functional groups such as α,β-unsaturated carbonyls, epoxy, and aldehyde. These functionalities are mostly formed through radical mechanism of including addition of oxygen or rearrangement of double bonds. It has been suggested that covalent binding of these electrophilic groups to the protein targets provides signal accumulation over time. This phenomenon has been named the covalent advantage. Thus, the full activation of the receptor can be achieved with a low concentration of oxidized lipids. Many targets have been identified as a target for LPO products such as
57-61 Cytochrome c oxidase, Keap1 protein, Hsp70, Hsp90, ANT and ATP synthase. The
key to these modifications is the presence of nucleophilic amino acids such as cysteine,
histidine and lysine that are available in most of the signaling proteins.62 Biological
22
activity of these molecules confirms that downstream of ROS/RNS results in a more specific impact in regulating cell signaling.
Since fatty acids are one of the major lipid groups that play a role as a source of energy, targets that are related to nutrient sensing were the first to be studied. In the following paragraphs, GPCR and PPAR are presented as examples of targets interacting with oxidized lipids. 63,64
1. 9.1 Oxidized lipid and G protein-coupled receptors (GPCRs)
Like many other oxidized lipids derived from enzymatic reactions such as prostanoids, oxidized free fatty acids derived from polyunsaturated fatty acids (PUFAs) are found to function by reversibly binding to cell G protein-coupled receptors (GPCRs).
These receptors are located on the surface of the cell, with seven domains spanning through the membrane. For example, in mast cells GPR132, as one of the members in proton-sensing GPCRs, has been found to interact with oxidized forms of fatty acids. It functions as a stress-inducible receptor that can recognize lipid overload and oxidative stress. 9-Hydroxyoctadecadienoic acid (9-HODE), an intermediate of LA peroxidation, has been identified as a ligand for this receptor. 65-67 In other studies, 4-HNE has demonstrated modulation of GPCR signaling through adduct formation directly with G
68,69 proteins such as Gαq/11 and RGS4.
23
COOH
LA
lipid peroxidation COOH HO 9-HODE
OH GPCR O
4-HNE
Mast cell
Muscrarinic receptor G protein G protein
Brain Cell
Figure1.6 Lipid peroxidation products and their signaling through GPCR
1. 9.2 Oxidized lipid and peroxisome-proliferator-activated receptors
(PPARs)
Among the intracellular targets, the lipid molecules are capable of promoting
signals via peroxisome-proliferator-activated receptors (PPARs) pathways in both
reversible and irreversible fashions. 70 The PPAR family of nuclear receptors is serving as
a master regulator of cellular differentiation, development and metabolism. Upon the binding of oxidized lipids to these receptors, their downstream genes are up-regulated.
PPARs also regulate the expression of other genes through interaction with coregulators
of gene expression in other gene machineries. (Section 3.2)
24
Cytoplasm
Lipid peroxidation
Co-activators
PPAR RXR9cRA
PPRE
Nucleus
Figure1.7 Lipid peroxidation products interact with PPAR and activate their gene expression machinery.
25
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30
Chapter 2 Development of novel synthetic routes to the
epoxyketooctadecenoic acids (EKODEs)
2.1 Introduction
Since the discovery of prostaglandins in the early 1930s by Ulf von Euler and M.
W. Goldblatt, scientists have been struggling for decades to define the mechanism of
action of oxidized lipids.1 In fact, advancements in instrumental methods such as mass
spectrometry and isotope labeling have provided great advantages for researchers to
isolate different oxidized lipids and characterize their structures in the 1960s.2
The broad and potent biological actions of these molecules have driven the
scientific community to investigate them more thoroughly. Further, access to knockout
mouse models for disruption of enzymes involved in lipid biosynthesis pathways and the
availability of genes for production of plausible sites of action have made these
discoveries possible.3
Among different research groups, Sune Bergström, Beng I. Samuelsson and John
R. Vane received the Nobel Prize in physiology or medicine in 1982 for their
“discoveries concerning prostaglandins and related biologically active substances”.
Accordingly, pioneering chemists in total synthesis such as Robert B. Woodward, Gilbert
Stork and Elias J. Corey have shown great interest in developing syntheses of these bioactive lipids.
Although the lipidologists in the field primarily focused on oxidized lipids formed
through enzymatic reactions, non-enzymatic products also became important to the field
in the 1960s. In fact, collaborative work between L. Jackson Roberts II and Robert G.
Salomon resulted in the discovery of lipid peroxidation products such as isoprostane and
31
isolevuglandin. In fact, non-enzymatically produced LPO products have proven to be as important as the enzymatic products in cell signaling.4
The general interest in the field of LPO is focused on biological target modifications with end products of LPO such as 4-HNE and 4-ONE (section1.3). Indeed, a systems analysis conducted by the Lawerence J. Marnett lab yielded interesting results, showing these molecules modulating biological networks. Likewise, his group revealed
4-HNE acts on different targets through the modulation of gene expression or directly on the protein targets.5-7 Recently in our laboratory detailed mechanistic studies showed that
4-HNE modulates the production of nitrogen oxide (NO.) in a concentration-dependent
manner in a macrophage culture cell line. In other words, 4-HNE acts as a second messenger in maintaining normal cell physiology.8
Currently in our laboratory, we are trying to look at lipid peroxidation products of
linoleic acid in a general prospective. In fact, we are examining the entire spectrum of
possible oxidized linoleic acid through lipid peroxidation as potential signaling
molecules. Linoleic acid (LA) is often the most prevalent form of PUFAs present inside
the cell membrane, and the 1,4-diene pattern makes it more vulnerable to oxidation than
other fatty acids.9 As a result, during oxidative stress many products of peroxidation are
derived from linoleic acids. Once ROS / RNS trigger the reaction by abstracting an
electron from LA, then molecule of oxygen adds to the carbon radical to generate
hydrorperoxy intermediate. Following the rearrangement or fragmentation, two classes of
products are formed: (1). Oxidative cleavage products (OCPs) which consist mainly of
the end products of peroxidation, such as 4-hydroxynonenal (4-HNE), and 4-oxononenal
32
(4-ONE). (2). Oxidative addition products (OAPs) resulting from oxygenation of radical intermediates along with rearrangement in double bonds.10 (Figure 2.1)
O O OH COOH COOH O EKODE OH DHKODE O O COOH COOH OH KODDE HKODE
ROS/RNS
COOH
LA
ROS/RNS
OH OH O O COOH
4-HNE HODA O O O O COOH 4-ONE DODE
COOH O O Octenal OUEA Figure 2.1 Linoleic acid oxidation through non-enzymatic reactions
Among these varieties of scaffolds of linoleic acid metabolites, epoxyketooctadecenoic acids (EKODEs) specifically caught our attention. This family of metabolites has been proven to exist in vivo and is involved in pathological states associated with oxidative stress such as asthma, obesity and hypertension.11-13 More detailed physiological studies showed that EKODEs change corticosterone and
33
aldosterone production levels. Evidence points to the latter being conducted through
modulation of Ca2+ concentration inside the cell.14 In addition, these oxidized lipids have
shown antioxidant effects through the Nrf2-Keap1 signaling pathway.15
The interesting physiological activities of EKODEs led us to consider a robust
synthesis for these scaffolds in a broader picture. Likewise, identifying methods of
synthesis for each core can be considered an innovative approach to gain access to an
array of metabolites and their modified forms.
2.2 EKODE structure and nomenclature
There are two series of EKODEs with different arrangements of carbonyl, epoxy and olefin functional groups. Isomers with C=C in the middle, and the others with C=O in the middle were assigned as EKODE-(E)-I and EKODE-(E)-II, respectively. Because the new C=C bonds form in free radical steps, they are more likely to form the trans (E) stereochemistry in the final products of EKODE. Epoxy groups are formed from hydroperoxy intermediates which can lead to both “trans / cis” epoxy, named with the same designation (Scheme 2.2, Scheme 2.7). In addition, “a” and “b” were used to define proximity of epoxy to the carboxylic acid or methyl terminals, respectively. Four isomers for each EKODE-I and EKODE-II can be proposed; however, the previous studies showed the cis-IIa / IIb are not formed (small amount) under peroxidation conditions.16
O O O
R1 R2 R1 R2 R1 R2 O O O trans-IIa/IIb trans-Ia/Ib cis-Ia/Ib
= R1 or R2 C5H11 = R or R (CH2)7COOH 2 1
Figure 2.2 EKODE I and II structures and nomenclature
34
2.3. EKODE-(E)-I family
R2 R1 R1=-(CH2)7COOH R2=-C5H11 LA radicals,
radicals , 2 O O 2
R2 O(O)H O(O)H R1
R1 R2 10E,12Z,9H(p)ODE 9Z,11E,13H(p)ODE
R R R2 R1 2 1 O O O O R R2 1 O2 O2 R2 R1 O O cis-EKODE-(E)-Ia cis-EKODE-(E)-Ib OOH OOH
R R R2 R1 2 O 1 O
O O
R R R2 R1 2 O 1 O trans-EKODE-(E)-Ia trans-EKODE-(E)-Ib Scheme 2.1 Non-enzymatic formation of EKODE-I family
The postulated non-enzymatic formation of both cis- and trans-EKODE-(E)-Ia and -b is summarized in Scheme 2.1. The first step involves the conversion of linoleic acid (LA) to 13 or 9-hydroperoxy (HpODE) intermediates, which is followed by rearrangement to a trans-epoxy and double bonds, producing a second site for oxygenation at the C-10,C-9 positions for Ia and Ib, respectively.
35
The hydroperoxy intermediate undergoes elimination to afford ketone as trans-
EKODE-(E)-Ia and -b. Similarly, if epoxy group is produced as cis stereoisomers, the
intermediates eventually lead to cis-EKODE-(E)-Ia and -b shown in dashed arrows in
Scheme2.1 In the next section, we illustrate a methodology to target total synthesis of
epoxy α, β-unsaturated ketone in the EKODE-(E)-I family of oxidized linoleic acid metabolites.
2.3.1 Retrosynthetic analysis of trans-EKODE-(E)- I
The retrosynthetic analysis for trans-EKODE-(E)-I family is outlined in Scheme
2.2 The trans-EKODE-(E)-I can be obtained from two smaller fragments of trans-epoxy aldehyde and the phosphorane through a Wittig reaction.
O
R1=-(CH2)7COOH R1,2 O R1,2 R2=-C5H11 Wittig olefination
O O Ph P R O + 3 R O 1,2O R Cl 1,2 1,2 R1,2 trans-epoxy aldehyde phosphorane
Wittig olefination
O O R1,2 HO R 1,2
Scheme2.2 Retrosynthesis analysis of trans-EKODE-(E)-I
In order to make the phosphorane; treatment of the carboxylic acid with oxalyl
chloride gave the acid chloride, which was reacted with methyltriphenylphosphorane,
generated by reaction of methyltriphenylphosphonium bromide and stoichiometric
amount of n-butyllithium at -78 oC, to give the phosphorane. For synthesis of the trans-
36
epoxy aldehyde moiety, starting from an aliphatic aldehyde, a one-step Wittig reaction with (triphenylphosphoranylidene)acetaldehyde gave α, β-unsaturated aldehyde, epoxidation of which, with alkaline H2O2, provided the trans-epoxy aldehyde. Finally, a
one-step Wittig reaction afforded the olefin.
2.3.2 Synthesis of trans-EKODE-(E)- Ia
The synthetic procedure for trans-EKODE-(E)-Ia was based on the preparation of trans-epoxy aldehyde (blue) and the phosphorane 7 (red).
In order to make the trans-epoxy aldehyde moiety, ozonolysis of commercially available methyl oleate gave methyl 9-oxononanoate 2 in 75 % yield, which was purified via distillation. A Wittig reaction of 2 with (triphenylphosphoranylidene)acetaldehyde gave methyl (E)-11-oxoundec-9-enoate 4 in a 57 % yield. Next, epoxidation of the α, β- unsaturated aldehyde with alkaline H2O2 afforded the trans-epoxide 5 in a 38 % yield.
In order to make the phosphorene 7, treatment of hexanoic acid with oxalyl chloride gave hexanoyl chloride in a 91 % yield. The latter was reacted with methyltriphenylphosphorane, generated by reaction of methyltriphenylphosphonium bromide and stoichiometric amount of n-butyllithium at -78 oC , to give 1-(triphenyl-λ5-
phosphanylidene) heptan-2-one 7.
37
O Ph P O O 3 3 O H2O2 Methyl oleate MeO O 3 MeO O 5 5 Zn CHCl3, reflux, 48 h NaHCO3, rt, 3h 1 (C19H36O2) 2 75 % 4 57 %
O O 1. (COCl)2, CH2Cl2, reflux,2h Ph3P HOOC MeO O + 5 O PPh Br n -78 o to rt 38 % 5 7 41 % 2. CH3 3 / -BuLi, C 6 Wittig reaction o CH2Cl2, 0 C, 4h
O O
MeO 5 O 47 % 8
Water / Acetone, pH 7 PPL 4h
O O
HO 5 O 93 % trans-EKODE-(E)-Ia Scheme 2.3 Synthetic route for trans-EKODE-(E)-Ia
The reaction of 7 (red) and trans-epoxy aldehyde 5 (blue) moieties at 0 oC
provided the methyl ester form of trans-EKODE-Ia 8, which was hydrolyzed with porcine pancreas lipase (PPL) to afford the final product trans-EKODE-(E)-Ia in a quantitative yield.
2.3.3 Synthesis of trans-EKODE-(E)-Ib
The synthetic procedure for trans-EKODE-(E)-Ib was based on the preparation of trans-epoxy aldehyde (pink) and the phosphorane 10 (green).
In order to make the phosphorane, methyl 9-oxononanoate 9 obtained from ozonolysis was transformed to its corresponding carboxylic acid in a 95 % yield through Pinnick oxidation. Next, treatment of 9 with oxalyl chloride gave methyl 9-chloro-9-
38
oxononanoate in a 93 % yield. The latter was reacted with methyltriphenylphosphorane,
generated by reaction of methyltriphenylphosphonium bromide and stoichiometric
amount of n-butyllithium at -78 oC , to give methyl 9-oxo-10-(triphenyl-λ5-
phosphanylidene) decanoate 10. In order to produce the trans-epoxy aldehyde 12,
epoxidation of commercially available (E)-oct-2-enal 11 with alkaline H2O2 gave 12 in a
63 % yield.
A reaction of 10 (green) and trans-epoxy aldehyde 12 (pink) at 0 oC provided
methyl ester 13 ,which was hydrolyzed with porcine pancreas lipase (PPL) to afford the
final product trans-EKODE-(E)-Ia in a quantitative yield.
O O COOH1. (COCl)2, CH2Cl2, reflux, 2h NaClO2 MeO MeO O o 5 5 H O ,10 2h n o 2 2 2 C, 95 % 9 2. CH3PPh3Br / -BuLi, -78 C to rt
O O H O PPh3 2 2 MeO +O O O 5 NaHCO 50 % 10 12 63 % 3 11
Wittig reaction o CH2Cl2, 0 C, 4h
O O
MeO 5 O 56 % 13
Water / Acetone,pH 7 PPL 4 h O O
HO 5 O Scheme 2.4 Synthetic route for trans-EKODE-(E)-Ib
39
2.3.4 Retrosynthetic analysis of cis-EKODE-(E)- Ib
cis-EKODE-(E)-I was prepared using a strategy similar to that for the trans analogues, although the cis-epoxy aldehyde was made through a completely different route. Starting from propargyl alcohol, alkylation of the terminal alkyne with bromopentane affords the acetylenic alcohol, which can be hydrogenated over Lindlar catalyst to afford a cis-olefin. Next, epoxidation provides the substrate for further oxidation with PCC in order to produce the cis-epoxy aldehyde.
O HO HO O O
HO HO
Scheme 2.5 Retrosynthesis analysis of cis-epoxy aldehyde
2.3.5 Synthesis of cis-EKODE-(E)- Ib
The synthetic procedure for trans-EKODE-(E)-Ib followed from the preparation of cis-epoxy aldehyde 18 (orange) and the phosphorane 10 (green).
The ylide 10 was made as described in section 2.2.3. In order to make the cis-epoxy aldehyde moiety, propargyl alcohol 14 was reacted with 2 equivalents of n-butyllithium at
-78 oC and then treated with n-bromopentane to give oct-2-yn-1-ol 15 in a 67 % yield.
Subsequent hydrogenation of the acetylenic alcohol intermediate over Lindlar catalyst
afforded (Z)-oct-2-en-1-ol 16 in a 92 % yield. Next, epoxidation with m-
chloroperoxybenzoic acid (m-CPBA) provided cis-(3-pentyloxiran-2-yl)methanol 17 with
40 a 95 % yield. Further oxidation with PCC gave cis-3-pentyloxirane-2-carbaldehyde 18, which is the required cis-epoxy aldehyde moiety.
HO n-bromopentane H2 HO n Lindlar catalyst, 8 h 14 H -BuLi, HMPA, THF 15 67 %
m-CPBA HO HO O CH2Cl2, rt, 2h 16 92 % 17 95 %
PCC CH2Cl2, rt, 2h
O O PPh3 + MeO O 5 O 50 % 10 18 55 %
Wittig reaction o CH2Cl2, 0 C, 4 h
O O
MeO 5 51 % 19 O
Water / Acetone, pH 7 PPL 4 h
O O
HO 5 O 93 % cis-EKODE-(E)-Ib
Scheme 2.6. Synthetic route for cis-EKODE-(E)-Ib
41
Reaction of 10 (green) with cis-epoxy aldehyde 18 (orange) at 0 oC provided the
methyl ester of cis-EKODE-(E)-Ib 19, which was hydrolyzed with porcine pancreas lipase (PPL) to afford the the final product cis-EKODE-(E)-Ib in a quantitative yield.
2.4 EKODE-(E)-II family
R R 2 O(O)H O2 R2 R1 O2 O(O)H 1
R1 R2 10E,12Z,9H(p)ODE Linoleic acid(LA) 9Z,11E,13H(p)ODE
R1=-(CH2)7COOH R2=-C5H11 R R R R 2 O 1 2 O 1
O2 O2
OOH OOH
R R R2 R1 2 O 1 O
O O
R2 R1 R R O 2 O 1 trans-EKODE-(E)-IIb trans-EKODE-(E)-IIa
Scheme 2.7 Non-enzymatic formation of EKODE-II family
The postulated non-enzymatic formation of trans-EKODE-(E)-IIa and-b is summarized in Scheme 2.7. The first step involves the conversion of LA to 13 or 9- hydroperoxy (HpODE) intermediates. Then, rearrangement to an epoxy functionality leads to the second site for oxygenation to a new hydroperoxide at the C-10 position, which is eventually undergoes elimination to afford ketone as trans-EKODE-(E)-IIa and- b.
42
The previous synthetic strategy for making specific arrangement of functional groups in trans-EKODE-(E)-II family involved formation of β-hydroxyenone
intermediate (X) (Scheme 2.8). Starting from an α, β-unsaturated ketone, aldol addition
o of aldehyde (R2CHO) using LDA / THF at -78 C is extremely low yielding. In fact
formation of side-products such as, tetrahydropyran through 6-endo-trig oxa-Michael
cyclization, during aldol reaction or purification on silica gel, make it an inefficient
synthetic route (Scheme 2.8i). 16
O O O OH R2 i. R R1 R2 1 aldol reaction X
O O NOH N R R2 red. 2 X ii. R R R1 1 (3+2) cycloaddition 1
O O O NOH O N O R R red. P 2 2 iii. (RO)2P (RO)2 (RO)2P
O O O OH R1 (RO)2P X R2 Scheme 2.8 Different synthetic strategies in order to make intermediate X.
There are a number of other approaches to avoid aldol addition while generating
intermediate X. For example, the second strategy (Scheme 2.8ii) involves taking
advantage of the reduction of a Δ2-isoxazoline in order to form the β-hydroxyenone. The key step for synthesis through this route is the isoxazoline ring formation.17 Formation of
the five-membered ring through 1,3-dipolar cycloaddition reaction stuffers from poor
regioselecvtivity and makes this approach challenging. In addition, reduction of Δ2-
43
isoxazoline using a variety of methods does not exhibit acceptable regioselectivity, and
there is a noticeable amount of reduced olefin in the final mixture.Finally, a third
approach which would avoid having a double bond in the starting material by using HWE
reagent, also experienced problems with reducing conditions mostly because of side
reactions such as elimination of the hydroxyl group (Scheme 2.8ii).
The previous synthetic routes have several disadvantages and problems: being
laborious, time-consuming, and complicated in terms of number of undesired products.16
Consequently, we sought to exploit a much more efficient tactic to avoid generation of
highly reactive β-hydroxyenone intermediate X.
In the next section, a new procedure has been developed in our laboratory based
on bifunctional conjunctive ylides to overcome the difficulties listed. Specifically, we
describe new, highly chemoselective, sequential reactions of sulfonium and Wittig ylides
with two different aldehydes that can be used for rapid construction of the trans-EKODE-
( E)-II family.
2.4.1 Retrosynthetic analysis of trans-EKODE-(E)-II
Structurally, this family of EKODEs possesses flanking α, β-unsaturated and epoxy functionalities centered on a ketone group. The synthetic method relies on developing a bifunctional intermediate containing two reactive ylides that can be used for generation of epoxy and olefin functionality in a sequential order. Our retrosynthetic analysis for the synthesis of the trans-EKODE-(E)-II family is outlined in Scheme 2.9, and starts from a bifunctional conjunctive ylide. Addition of the first aldehyde through
Johnson-Corey-Chaykovsky reaction produces the triphenylposphine epoxy ketone.
44
Subsequent Wittig olefination with a second aldehyde provided α,β-unsaturated keto
epoxide functionality.
O Wittig olefination O Ph3P R R R 2 O 1 O 1
Johnson–Corey–Chaykovsky reaction
O
Ph3P S(CH3)2
Bifunctional conjunctive ylide Scheme 2.9 Retrosynthesis analysis of trans-Ekode-(E)-II
2.4.2 Synthesis of bifunctional conjunctive ylide
Scheme 2.10 shows the detailed synthesis of bifunctional conjunctive ylide 22.
The synthesis of 22 began with commercially available 1,3-dichloroacetone. Halides in
1,3-dichloroacetone were replaced (Finkelstein reaction, 90 %) by bromide,18 and
substitution of one of the bromides with triphenylphosphine (90 %) followed by base
treatment that provided the Wittig ylide.18 Addition of dimethylsulfide in the presence of sodium tetrafluoroborate afforded the bifunctional conjunctive ylide (77 %). The overall yield was 35 %.
45
O O
Cl Cl 15 eq LiBr Br Br 1. 1.1eq PPh3 / Toluene, rt, overnight
19 Acetone, rt, 3days 20 2. NaHCO3, Methanol/water 90 %
O O Ph P Ph P - 3 Br 5 eq S(CH3)2, 5eq NaBF4 3 S(CH3)2 BF4 Acetone, rt, 24 hrs 21 22 Bifunctional conjunctive ylide 50 % 77 % Scheme 2.10 Synthetic rout for bifunctional conjunctive ylide
A literature review shows that the application of bis-ylides has been known for over a half century. However, their application has been limited to the synthesis of symmetric olefins via Wittig and related types of olefination reactions. 19,20 Recently,
Nagorny et al., developed a series of asymmetric bifunctional ylides in order to make
asymmetric polyene. 21
In this novel approach, the bis-ylides used in the coupling possess different
reactivities toward electrophiles. Appel et al. showed that a triphenylphosphonium ylide
has lower kinetic reactivity compared to the sulfonium version. 22 In fact, in a
phosphonium ylide the carbanion is more stabilized by the adjacent heteroatom than the
corresponding sulfonium derivative. The main cause in the case of phosphonium has been
attributed to the electron withdrawing effect through pπ-dπ which is pulling away
electrons from carbanion. Consequently, this phenomenon exhibits a substantial impact
on the reactivity toward aldehyde electrophiles. This difference in reactivity has enabled
the development of a selective sequence of reactions with respect to the aldehydes.
46
So far there have been only a few examples in which the difference in reaction
rate between ylide reagents has been used. 23,24 However, their application in the total synthesis of natural products has not yet been explored.
2.4.3 Synthesis of trans-EKODE-(E)-II
In the first step, the bifunctional conjunctive ylide was treated with sodium hydride and then with aldehyde (R1CHO) to form an epoxide. Formation of the epoxide
with n-hexanal 24 occurs much faster at lower temperature (at 0 oC for 3 h). However
methyl 9-oxononanoate 2 needs to be warmed to room temperature and requires a longer
time to complete the reaction (rt for 8 h). The yields for the reactions of n-hexanal and
methyl 9-oxononanoate were 72 and 58 percent, respectively. The olefin was formed
from the epoxide intermediate after treatment with the second aldehyde (R2CHO); n-
hexanal and methyl 9-oxononanoate gave yields of 76 and 62 percent, respectively. The
olefination reaction gave primarily trans stereoisomer. Finally, hydrolysis with porcine pancreas lipase (PPL) gave rise to the final product. 25 Starting from intermediate 22, the
overall yield was 60 %.
O O O 1 eq R1CHO Ph3P 1 eq R2CHO Ph3P S(CH3)2 R R2 R1 O 1 O 22
Bifunctional conjunctive ylide Epoxide formation Olefin formation Scheme 2.11 General strategy for synthesis of trans-EKODE-(E)-II family
The advantage of a bifunctional conjunctive ylide is that it represents two
different types of ylides such as sulfonium and phosphonium in the same molecule.
Consequently, this intermediate provides an opportunity to perform corresponding types of reactions with aldehydes, such as Corey-epoxidation and Wittig olefination. The presented structure of the trans-EKODE-(E)-II family in Figure 2.1 shows a palindromic
47
sequence of functional groups. In other words, by using two different aldehydes and
changing the order of addition to the ylides, both products can be obtained. The synthetic
route for trans-EKODE-(E)-IIa is presented in Scheme 2.12. The reaction of aldehyde 2
with the sulfonium ylide forms epoxy intermediate 23 (58 %), after which the
phosphonium ylide reacts with n-hexanal 24 and forms the olefin 25 (76
%).
O
1eq O OMe ,1eq NaH O O O O 5 1eq Ph3P Ph P 2 OMe 24 3 S(CH3)2 5 o O 22 Acetonitrile, -30 C to rt, overnight 23 Benzene, reflux, 48 h
O O O O PPL OMe OH O 5 Acetone, water, 3 h O 5 trans-EKODE-(E)-IIa 25
Scheme2.12 Synthetic route for trans-EKODE-(E)-IIa
Similarly, the synthetic route for trans-EKODE-(E)-IIa is presented in Scheme
2.12. The reaction of aldehyde 24 with sulfonium ylide formed an epoxy intermediate 26
(72%), after which the phosphonium ylide reacted with n-hexanal 24 and formed the olefin 27 (62 %).
48
O O O O NaH 1eq O OMe 1eq ,1eq Ph P 5 3 2 Ph3P S(CH3)2 24 o O 22 Acetonitrile, -30 C to rt, 3 h 26 Benzene, reflux, 48 h
O O O O PPL HO MeO 5 O 5 O Acetone, water, 3 h 27 trans-EKODE-(E)-IIb Scheme 2.13 Synthetic route for trans-EKODE-(E)-IIa an b
The described method was applied to the synthesis of the trans-EKODE-(E)-II family. Our synthesis commenced from bifunctional conjunctive ylide with the addition of two known aldehydes. Finally, hydrolysis with porcine pancreas lipase (PPL) gave rise to the final product in quantitative yield.25 By changing the order of addition for two
aldehydes, both trans-EKODE-(E)-IIa and -b were obtained. Starting from intermediate
22, the overall yield was 41 % for trans-EKODE-(E)-IIa and –b.
2.4.4 Divergent synthesis for the varied library of trans-EKODE-(E)-II
Since our initial intention was studying physiological roles of the trans-EKODE-
(E)-II family, we envisioned that the synthesis of modified alkyne-terminal and deuterated-labeled lipids could be useful for our purpose.
2.4.4.1 Synthesis of alkyne-end aldehyde
To make the alkyne-end aldehyde, propargyl alcohol 14 was reacted with 2eq amount of n-butyllithium at -78 oC and treated with n-bromopropane to give hex-2-yn-1-
ol 28 (57 % ). Next, the alkyne-zipper reaction provided by a strong base moved the triple
bond to the terminal end of the molecule to form terminal alkyne in product 29.
Oxidation of primary alcohol with PCC provided an aldehyde 30 (68 %).
49
n-BuLi, 1-bromopropane HO Li, , 70 o 1h HO (CH2)3(NH2)2 C, o H THF / HMPA -78 C to rt / overnight 28 t-BuOK, rt, 3h 14
PCC HO O DCM, 3h 30 29
Scheme 2.14 Synthetic route for alkyne-end aldehyde
2.4.4.2. Synthesis of deuterated labeled aldehyde
To make the deuterated-labeled aldehyde, d11-carboxylic acid 31 was reduced with LiAlH4, followed by oxidation of alcohol 32 with PCC to obtain an aldehyde.
D D D D D D D D D D D D PCC LiAlH4,ether, 2h HOH C HOOC 2 CD3 O CD3 CD3 D D D D CH2Cl2, 3h D D D D D D D D
32 33 31
acid Hexanoic-d11
Scheme 2.15 Synthetic route for deuterated-labeled aldehyde
Bifunctional conjunctive ylide was treated with sodium hydride and then aldehyde
(R1CHO) to form an epoxide. Yields are between. 58-72%.
50
O - O Ph3P 1eq R1CHO, 1eq NaH S(CH3)2 BF4 Ph3P o R Acetonitrile, -30 C to rt, overnight O 1 Bifunctional conjuctive ylide
entry aldehyde product yield(%)*
O O O 1 OMe Ph3P 58 O 5 OMe O 5 2 23 O 2 Ph3P O 72 O 24 26
O Ph P 3 3 67 O O 34 30
D D D D O D D D D 4 Ph3P 70 O CD3 CD D O 3 D D D D D D D
33 35
Table 2.1 Reaction of sulfonium ylide and aldehydes, * yields were calculated based of amount obtained after purification
Epoxide intermediate was treated with aldehyde (R2CHO) and formed the olefin.
This reaction gave primarily trans stereoisomer. The yields were between 55- 76 %.
51
O O 1eq R CHO Ph3P 2 R R2 R1 O 1 Benzene, reflux, 48h O
entry substrate aldehyde product yield(%)*
O O 1 23 O O 76 24 O 5 25 O O
O O 71 2 23 30 O 5 36
D D D D D D D D O O 63 D C O O CD3 3 O 5 3 23 D D D D D D D D 37 33 O O O 62 O O 5 OMe 27 4 26 O 5 2 O O O O 5 55 OMe O 5 34 O 5 38 2 O O O D D D D D C O OMe 3 5 59 6 35 O O 5 D D D D 39 2
Table 2.2 Reaction of phosphonium ylide and aldehydes.* yields were calculated based of amount obtained after purification
In summary, we have described a divergent synthetic route to gain access to the
library of the trans-EKODE-(E)-II family and their alkyne-end and deuterium-labeled homologues. The route we present allowed us to prepare gram quantities of these molecules. Such a strategy might also be useful for the rapid generation of other lipid metabolites derived from other PUFAs with a similar combination of functionalities that are valuable as biologically active molecules.
52
2.5 Experimental
2.5.1 General experimental details
All reactions were performed in oven-dried glassware, under an argon atmosphere
with rigid exclusion of moisture from reagents and solvents. Hexanoic-d11 acid (98 atom
% 2H) and PPL (porcine pancreas lipase, Type II) were purchased from Sigma-Aldrich.
All other reagents were used as supplied by chemical manufacturers. The exact molarity of n-butyllithium was determined each time by titration prior to use with 2-propanol,
Et2O at 0 °C and 1,10-phenanthroline as the indicator. Tetrahydrofuran was distilled from a purple solution of sodium benzophenone ketyl. All other solvents were used as purchased.
Ozonolysis was performed on the ozone generator lab series (L21) (Pacific
Ozone), using industrial grade compressed oxygen (purity 99.5 %). Liquid
chromatography was performed using forced air-flow (flash chromatography) on silica
gel (230- 400 mesh), by eluting solvent (reported as V: V ratio mixture). Analytical thin
layer chromatography (TLC) was performed on Partisil K6F Silica Gel 60A 0.25 mm
plates with fluorescent indicator. Visualization of the developed chromatogram was
accomplished with UV light (254 nm) and stained with either ethanolic phosphomolybdic
acid (PMA) or ceric ammonium molybdate. 1H and 13C NMR spectra were either
recorded on a Varian Inova spectrometer operating at 400 MHz and 100 MHz for the 1H and 13C respectively or Bruker Ascend Avance III HDTM operating at 500 MHz and 125
MHz for the 1H and 13C respectively (Department of Chemistry, Case Western Reserve
University). The internal references were CDCl3 (δ 7.26) and CD3OD (δ 3.31) for 1H and
(δ 77.36) and CD3OD (δ 49.00) for 13C spectra, respectively. NMR data are presented in
53
the following order: chemical shift, peak multiplicity (b = broad, s = singlet, d = doublet,
t = triplet, q = quartet, m = multiplet, dd = doublet of doublet, ddd = doublet of doublet of
doublet, ddt = doublet of doublet of triplet, dq = doublet of quatrate, dm = doublet of
multiplet, br = broad), coupling constant (in Hz). Mass spectra were acquired on a
Thermo Scientific LTQ-FT hybrid mass spectrometer (Rieveschl Laboratories for Mass
Spectrometry, University of Cincinnati) using electrospray ionization (positive mode).
54
2.5.2 Syntheses
Methyl 9-oxononanoate (2) (C10H18O3)
COOMe O
An oven-dried 0.5-L, two-necked round-bottomed flask was equipped with a mechanical stirrer. The flask was charged with methyl oleate 1 (20.8 g, 70 mmol) in dichloromethane (250 mL) and methanol (100 mL). The vessel was cooled to -78 °C and ozone was bubbled through the solution using a Pasteur pipet until a persistent dark blue
color appeared. The solution was purged with argon gas until the color dissipated, and
then the cold bath was removed. The ozonide was decomposed by adding glacial acetic
acid (25ml) in one portion, followed by powdered Zn (9 g, 0.14 mol) in small portions to
control heat evolution. The mixture continued to stir for an additional half an hour and
then filtered through Celite to remove unreacted Zn in the mixture. Water (100 mL) was
added to the mixture to prevent acetal formation. The solution was concentrated to about
one-half the initial volume in vacuo and slowly added to saturated NaHCO3 (150 mL).
The mixture was extracted with CH2Cl2 (3 × 200 mL). The organic phases were dried and
concentrated. The crude product was distilled to produce nonanal (bp 44 oC, 1 Torr) and
9.82 (75 %) of methyl 9-oxononanoate 2 (bp 103 oC / 1 Torr) as a colorless oil. (lit.26 bp
105-108 oC / 1 Torr)(Caution: Ozone is highly toxic and can react explosively with
numerous oxidizable materials.) Identity and purity of the product were confirmed by 1H
26 1 NMR. Spectroscopic data were found to match lit. data. H NMR (400 MHz, CDCl3) δ
1.28-1.36 (m, 6H) 1.58-1.66 (m, 4), 2.42 (td, 2H, J = 7.2, 2.0 Hz), 3.66 (s, 3H), 9.76 (t,
3H, J = 2 Hz).
55
Methyl (E)-11-oxoundec-9-enoate (4) (C12H20O3)
COOMe O
An oven-dried 50-mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with methyl 9-oxononanoate 3 (1.90 g, 10.22 mmol) and (triphenylphosphoranylidene)acetaldehyde 3 (3.72 g, 12.26 mmol) in dry chloroform
(20 mL). The mixture was heated under reflux at 70 oC for 48 h. Next, water (30 mL) was
added to the mixture, which was extracted with CH2Cl2 (3 × 30 mL).The combined
organic phases were dried with anhydrous magnesium sulfate and concentrated under
reduced pressure. Purification by column chromatography (Hexanes: ether / 80: 20)
afforded 1.23 g (57 %) of methyl (E)-11-oxoundec-9-enoate 4 as a yellow oil. Identity and purity of the product were confirmed by 1H NMR. Spectroscopic data were found to
1 match lit. data.16 H NMR (400 MHz, CDCl ) δ 1.28-1.39 (6H), 1.45-1.69 (4H), 2.28- 3
2.37(m, 4H), 3.67 (s, 3H), 6.08 (ddt, 1H, J = 15.6, 8.0, 1.6 Hz), 6.85 (dt, 1H, J = 15.6, 6.8
Hz), 9.50 (d, 1H, J = 8.0 Hz).
trans-methyl 8-(3-formyloxiran-2-yl)octanoate (5) ( C12H20O4)
COOMe O O
An oven-dried 50-mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with methyl (E)-11-oxoundec-9-enoate 4 (1.20 g, 5.66 mmol) in MeOH (15 mL). The flask was cooled in an ice-bath and NaHCO3 (570 mg,
6.78 mmol) was added. The reaction mixture was cooled between 0 °C, and H2O2
solution (1.7 mL, 16.7 mmol, 30% solution) was added dropwise via syringe. The
56
resulting mixture was vigorously stirred at ambient temperature for 3 h. The resulting
suspension was cooled between 0-5 °C in an ice bath and the excess hydrogen peroxide
was quenched with a saturated solution of sodium thiosulfate (2.0 mL dropwise). The
mixture was concentrated by rotary evaporation at room temperature. The aqueous phase
was extracted with EtOAc (3 × 10 mL) the combined organic phases dried with
anhydrous magnesium sulfate and concentrated under reduced pressure. Purification by
column chromatography (Hexanes: ether / 75: 25) afforded 488 mg (38 %) of trans-
methyl 8-(3-formyloxiran-2-yl)octanoate 5 as yellow oil. Identity and purity of the
product were confirmed by 1H NMR. Spectroscopic data were found to match lit. data.16
1 H NMR (400 MHz, CDCl3) δ 1.22-1.52 (8H), 1.52-1.70 (4H), 2.32 (t, 2H, J = 7.2 Hz),
3.15 (dd, 1H, J = 6.4 Hz and 2.0 Hz), 3.23 (td, 1H, J = 5.6 Hz and 2.0 Hz), 3.68 (s, 3H),
9.02 (d, 1H, J = 6.4 Hz).
5 1-(triphenyl-l λ -phosphanylidene)heptan-2-one (7) ( C25H27OP)
O Ph P 3 An oven-dried 50-mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with hexanoic acid 6 (2.09 g, 18 mmol). Then oxalyl
chloride (27 mmol) was added slowly via syringe under argon at room temperature. Once
evolution of gas had stopped (ca. 30 min), the mixture was heated under reflux for 2 h
and then cooled to room temperature. The excess oxalyl chloride was distilled at
atmospheric pressure and the remaining material was distilled to produce 2.20 g (91 %) hexanoyl chloride (bp 145 oC / 750 Torr) as a colorless oil. (lit.27 bp 151-153 oC / 1 Torr).
It was directly used for the next reaction.
57
Next, an oven-dried 100-mL, two-necked round-bottomed flask was equipped
with a magnetic stirring bar and sealed under argon with two rubber septa, one of which
contained a needle adapter, to an argon-inlet. The flask was charged with methyltriphenyl-phosphonium bromide (3.66 g, 10.26 mmol) in THF (20 mL). The solution was cooled in a dry ice-acetone bath at -78 °C. Then n-butyllithium (2.12 mL,
2.42 M in hexane, 5.13 mmol) was added, causing a red color to develop. The reaction
mixture continued to stir for an additional 1 h, whereupon hexanoyl chloride (690 mg,
5.13 mmol) was added dropwise slowly via syringe. The mixture was then allowed to stir
at ambient temperature for 1 h. The reaction mixture was neutralized with aqueous
saturated NH4Cl (10 mL) and evaporated the THF and resulted a viscous oil. The oil was extracted with EtOAc( 3 × 20 mL) and washed with NaOH (2 N, 50 mL).The combined organic phases were dried with anhydrous magnesium sulfate and concentrated under reduced pressure. Purification by column chromatography (DCM: Methanol / 98: 2) afforded 860 mg (45 %) of 1-(triphenyl-l λ5-phosphanylidene)heptan-2-one 7 as brown oil. Identity and purity of the product were confirmed by 1H NMR. 1H NMR (500 MHz,
2 CDCl3) δ 0.88 (bs, 3H,), 1.1-1.89 (m, 4H), 2.3 (bs, 2H), 3.72(d, 1H, JHP = 26.0 Hz), 7.42-
7.71 (m, 15H).
Methyl (E)-8-(3-(3-oxooct-1-en-1-yl)oxiran-2-yl)octanoate (8) ( C19H32O4)
O COOMe O
An oven-dried 50-mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with 1-(triphenyl-l λ5-phosphanylidene)heptan-2-one
58
7 (749mg, 2 mmol) and CH2Cl2 (10 mL). The resulting mixture was cooled to 0 °C in an ice bath. Then trans-methyl 8-(3-formyloxiran-2-yl)octanoate 5 (456mg, 2 mmol) was
added in CH2Cl2 (10 mL) via syringe. The reaction mixture continued to stir for an
additional 4 h at 0 °C. Next water (10 mL) was added to the mixture and extracted with
CH2Cl2 (3 × 20 mL). The combined organic phases were dried with anhydrous
magnesium sulfate and concentrated under reduced pressure. Purification by column
chromatography (Hexanes: ether / 85: 15) afforded 305 mg (47%) of methyl (E)-8-(3-(3-
oxooct-1-en-1-yl)oxiran-2-yl) octanoate 8 as a white oil. Identity and purity of the
product were confirmed by 1H NMR and 13CNMR. Spectroscopic data were found to
16 1 match lit. data. H NMR ( 500 MHz, CDCl3) δ 0.89 (t, 3H, J = 7.0 Hz), 1.22-1.69 (m,
18H), 2.30 (t, 2H, J = 7.5 Hz), 2.53 (t, 1H, J = 7.5 Hz), 2.89 (td, 1H, J = 5.5, 2.0 Hz),
3.20 (dd, 1H, J = 7.0, 2.0 Hz), 3.67 (s, 1H), 6.38 (d, 1H, J = 16.0 Hz), 6.51 (dd, 1H, J =
13 16.0, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 14.26, 22.79, 24.06, 25.20, 26.09, 29.31,
29.43, 29.46, 31.74, 32.22, 34.37, 41.00, 51.82, 56.97, 61.88, 131.69, 142.77, 174.60,
200.09.
General procedure: PPL hydrolysis
An oven-dried 25-mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with 0.1 mmol of the desired methyl ester EKODE-
(E) and PPL (220 mg) in acetone (1 ml) and phosphate buffer (pH 7, 5 mL, 0.2 M). The
mixture was stirred vigorously at ambient temperature for 4h and filtered through a pad
of Celite with EtOAc. The two phases separated and the organic phase was concentrated
59
under reduced pressure. Purification by column chromatography (Hexane: EtOAc / 50:
50) afforded the final product of carboxylic acids.
(E)-8-(3-(3-oxooct-1-en-1-yl)oxiran-2-yl)octanoic acid (trans-EKODE-(E)-Ia)
(C18H30O4)
O COOH O
Following the general procedure for PPL hydrolysis of methyl (E)-8-(3-(3- oxooct-1-en-1-yl)oxiran-2-yl)octanoate 8 afforded 29mg(93 %) of trans-EKODE-(E)-Ia as white solid. Identity and purity of the product were confirmed by 1H NMR and
13CNMR. Spectroscopic data were found to match lit. data.16 1H NMR ( 500 MHz,
CDCl3) δ 0.88 (t, 3H, J = 7.0 Hz), 1.23-1.69 (m, 18H), 2.35 (t, 2H, J = 7.5 Hz), 2.53 (t,
1H, J = 7.0 Hz), 2.91 (td, 1H, J = 5.5, 2.0 Hz), 3.21 (dd, 1H, J = 7.0, 2.0 Hz), 6.39 (d, 1H,
13 J = 15.5 Hz), 6.51 (dd, 1H, J = 15.5, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 14.26,
22.79, 24.07, 24.94, 26.08, 29.22, 29.41, 29.44, 31.75, 32.21, 34.09, 40.96, 56.99, 61.89,
131.70, 142.80, 180.50, 200.18.
9-methoxy-9-oxononanoic acid (9) (C10H18O4)
HOOC COOMe
An oven-dried 100-mL, two-necked round-bottomed flask containing a magnetic
stirring bar was equipped with an equalizing dropping funnel and an argon inlet. The flask was charged with methyl 9-oxononanoate 2 (2.8 g, 13.8 mmol) in acetonitrile (15 mL) and NaH2PO4 (480 mg) in water (6 mL) and H2O2 (1.74 mL, 17.1 mmol, 30 %). A
solution of NaClO2 (2.4 g, 21 mmol, 80 % purity) in water (20 mL) was added drop-wise
60
over 2 h to a stirred mixture while keeping the temperature at 10 oC with cooling. A small
amount (0.2 g) of Na2SO3 was added to destroy the unreacted HOCl and H2O2. The
mixture was acidified with 10 % aqueous HCl to pH 1 and concentrated by rotary
evaporation at room temperature. The aqueous phase was extracted with EtOAc (3 × 10
mL), the combined organic phases dried with anhydrous magnesium sulfate and
concentrated under reduced pressure afforded 2.7 g (95 %) of 9-methoxy-9-oxononanoic
acid 9 as a colorless oil. Identity and purity of the product were confirmed by 1H and 13C
28 1 NMR. Spectroscopic data were found to match lit data. H NMR (400 MHz, CDCl3) δ
1.25-1.38 (m, 6H), 1.56-1.70 (m, 4H), 2.3 (t, 2H, J = 17.6 Hz), 2.34 (t, 2H, J = 17.6 Hz),
13 3.66 (s, 3H); C NMR (100 MHz, CDCl3) δ 24.86, 25.13, 29.12, 29.16, 29.19, 34.33,
51.83, 174.68, 180.43.
Methyl 9-oxo-10-(triphenyl-l5-phosphanylidene)decanoate (10) (C29H33O3P)
O Ph P COOMe 3 An oven-dried 50-mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with 9-methoxy-9-oxononanoic acid 9 (2.43 g, 12 mmol). Then oxalyl chloride (18 mmol) was added slowly via syringe under argon at room temperature. Once evolution of gas had stopped (ca. 30 min), the mixture was heated under reflux for 2 h and then cooled to room temperature. The excess oxalyl chloride was distilled at atmospheric pressure and the remaining material was distilled to produce 2.50 mg (94 %) of methyl 9-chloro-9-oxononanoate (bp 125 oC / 1 Torr) as a
colorless oil. (lit.29 bp 125-127 oC / 1 Torr) Identity and purity of the product were
confirmed by 1H NMR. Spectroscopic data were found to match lit. data.16 1H NMR (500
61
MHz, CDCl3) δ 1.25-1.39 (m, 1H), 1.58-1.75 (m, 1H), 2.30 (t, 2H, J = 7.0 Hz), 2.88 (t,
2H, J = 7.0 Hz), 3.67 (s, 3H).
Next, an oven-dried 100-mL, two-necked round-bottomed flask was equipped
with a magnetic stirring bar and was sealed under argon with two rubber septa, one of
which contained a needle adapter, to an argon-inlet. The flask was charged with methyltriphenyl-phosphonium bromide (3.66 g, 10.26 mmol) in THF (20 mL). The solution was cooled in a dry ice-acetone bath at -78 °C. Then n-butyllithium (2.20 mL,
2.33 M in hexane, 10.26 mmol) was added, causing a red color to develop. The reaction mixture continued to stir for an additional 1 h, whereupon methyl 9-chloro-9- oxononanoate (1.13 g, 5.13 mmol) was added dropwise slowly via syringe, the mixture was then allowed to stir at ambient temperature for an additional1 h. The reaction mixture was neutralized with aqueous saturated NH4Cl (10 mL) and evaporated the THF,
resulting in a viscous oil. The oil was extracted with EtOAc (3 × 20 mL) and washed
with NaOH (2 N, 100 mL).The combined organic phases were dried with anhydrous
magnesium sulfate and concentrated under reduced pressure, affording 2.5 g (53 %) of
methyl 9-oxo-10-(triphenyl-l5-phosphanylidene)decanoate 10 as a brown oil. The
product was used as crude in its crude form in the Wittig reaction.
trans-3-pentyloxirane-2-carbaldehyde (12) (C8H14O2)
O O An oven-dried, 50-mL, round-bottomed flask was equipped with a magnetic stirring bar. The flask was charged with (E)-oct-2-enal 11 (714 mg, 5.66 mmol) in MeOH
(15 mL). The flask was cooled in an ice-bath and NaHCO3 (570 mg, 6.78 mmol) was
62
added. The reaction mixture was cooled to 0 °C, and H2O2 solution (1.7 mL, 16.7 mmol,
30 %solution) was added dropwise via syringe. The resulting mixture was vigorously stirred at ambient temperature for 1.5 h. The resulting suspension was cooled between 0-
5 °C in an ice bath and the excess hydrogen peroxide was quenched with a saturated
solution of sodium thiosulfate (2.0 mL dropwise). The mixture was concentrated by
rotary evaporation at room temperature. The aqueous phase was extracted with EtOAc (3
× 10 mL), and the combined organic phases were dried with anhydrous magnesium sulfate and concentrated under reduced pressure. Purification by column chromatography
(Hexanes: ether / 75: 25) afforded 510mg (63 %) of trans-3-pentyloxirane-2- carbaldehyde 12 as yellow oil. Identity and purity of the product were confirmed by 1H
and 13C NMR. Spectroscopic data were found to match lit. data.16 1H NMR (400 MHz,
CDCl3) δ 0.89 (t, 3H, J = 7.2 Hz), 1.27-1.70 (m, 8H), 3.13 (dd, 1H, J = 6.4, 2.0 Hz), 3.21-
3.24 (m, 1H), 9.01 (d, 1H, J = 6.4 Hz).
Methyl (E)-9-oxo-11-(3-pentyloxiran-2-yl)undec-10-enoate (13) (C19H32O4)
O O
MeO O
An oven-dried 50- mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with methyl 9-oxo-10-(triphenyl-l5-phosphanylidene) decanoate 10 (921 mg, 2 mmol) and CH2Cl2 (10 mL). The resulting mixture was cooled to 0 °C in an ice bath. Trans-3-pentyloxirane-2-carbaldehyde 5 (284 mg, 2 mmol) was added in CH2Cl2 (10 mL) via syringe. The reaction mixture continued to stir for an
additional4 h at 0 °C. Next, water (10 mL) was added to the mixture and extracted with
CH2Cl2 (3 × 20 mL). The combined organic phases were dried with anhydrous
63
magnesium sulfate and concentrated under reduced pressure. Purification by column
chromatography (Hexanes: ether / 85: 15) afforded 363 mg (56 %) of methyl (E)-9-oxo-
11-(3-pentyloxiran-2-yl)undec-10-enoate 13 as a yellow oil. Identity and purity of the product were confirmed by 1H and 13C NMR. Spectroscopic data were found to match lit.
16 1 data. H NMR (500 MHz, CDCl3) δ 0.89 (t, 3H, J = 7.0 Hz), 1.24-1.67 (m, 18H), 2.34
(t, 2H, J = 7.5 Hz), 2.53 (td, 1H, J = 7.0, 2.0 Hz), 2.91 (td, 1H, J = 7.0, 2.0 Hz), 3.21 (dd,
1H, J = 7.0, 2.0 Hz), 3.63 (s, 3H), 6.35 (d, 1H, J = 16.0 Hz), 6.45(dd, 1H, J = 15.5, 7.0
13 Hz); C NMR (125 MHz, CDCl3) δ 14.31, 22.87, 24.24, 25.20, 25.84, 29.27, 29.35(2C),
31.86, 32.24, 34.39, 40.90, 51.83, 56.99, 61.99, 131.62, 142.92, 174.63, 200.02.
(E)-9-oxo-11-(3-pentyloxiran-2-yl)undec-10-enoic acid (trans-EKODE-(E)-
Ib)(C18H30O4)
O O
HO O
Following the general procedure for PPL hydrolysis of methyl (E)-9-oxo-11-(3- pentyloxiran-2-yl)undec-10-enoate 13 afforded 28.0 mg (90 %) of trans-EKODE-(E)-Ib
as a yellow solid. Identity and purity of the product were confirmed by 1H and 13C NMR.
16 1 Spectroscopic data were found to match lit. data. H NMR (500 MHz, CDCl3) δ 0.89 (t,
3H, J = 7.0 Hz), 1.24-1.67 (m, 18H), 2.34 (t, 2H, J = 7.5 Hz), 2.53 (td, 1H, J = 7.0, 2.0
Hz), 2.91 (td, 1H, J = 7.0, 2.0 Hz), 3.21 (dd, 1H, J = 7.0, 2.0 Hz), 6.42(d, 1H, J = 15.5
13 Hz), 7.06(dd, 1H, J = 16.0, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 14.40, 22.87, 24.23,
24.92, 25.83, 29.16, 29.31(2C), 31.85, 32.23, 34.14, 40.86, 57.02, 62.00, 131.17, 142.93,
179.18, 200.04.
64
oct-2-yn-1-ol (15) (C8H14O)
HO
An oven-dried 500 mL, two-necked round-bottomed flask equipped with a magnetic stirring bar was sealed under argon with two rubber septa, one of which contained a needle adapter attached to an argon-filled balloon. The flask was charged with propargyl alcohol 14 (5.61 g, 0.1 mol), tetrahydrofurane (THF, 150 mL) and hexamethyl phosphoric triamide ( HMPA, 40 mL). The solution was cooled in a dry ice- acetone bath at -78 °C, and n-butyllithium (81.6 mL, 2.45 M in hexane, 0.2mol) was added slowly via syringe. The resulting solution was stirred for 5 min at -78 °C, after which it was slowly warmed to -30 °C. Following, 1-bromopentane (9.23 g, 75 mmol) was added slowly to the mixture via syringe. The temperature was slowly increased to ambient temperature overnight while the mixture stirred. The reaction mixture was neutralized with aqueous saturated NH4Cl (100 mL) and extracted with EtOAc (3 × 100 mL), the combined organic phases were dried with anhydrous sodium sulfate and
concentrated under reduced pressure. Purification by column chromatography (EtOAc:
Hexanes / 85: 15) afforded 6.30 g (67% yield) of oct-2-yn-1-ol 15 as a colorless oil.
Identity and purity of the product were confirmed by 1H and 13C NMR. Identity and
purity of the product were confirmed by 1H NMR. Spectroscopic data were found to
30 1 match lit. data. H NMR (400 MHz, CDCl3) δ 0.90 (t, 1H, J = 7.2 Hz), 1.28-1.39 (m,
4H), 1.51 (quint, 2H, J = 7.2 Hz), 2.21 (tt, 2H, J = 7.2, 2.0 Hz), 4.25 (t, 2H, J = 2.0 Hz);
13 C NMR (100 MHz, CDCl3) δ 14.29, 19.02, 22.53, 28.62, 31.36, 51.71, 78.56, 86.95.
65
(Z)-oct-2-en-1-ol (16) ( C8H16O)
HO
An oven-dried 1L round-bottomed flask was equipped with a magnetic stirring bar. The flask was charged with oct-2-yn-1-ol 15 (3.03 g, 24.0 mmol) and hexane (350
mL) under an argon atmosphere. To this solution, distilled quinoline (0.9 mL) and
Lindlar catalyst (0.36 g, 12% by weight) were added. The argon was evacuated by water aspirator and then filled with hydrogen from the balloon by adjusting the three-way valve. The reaction was monitored by 1H NMR and completed within 8 hours. The hydrogen atmosphere was replaced with argon and the mixture was filtered using a pad of
Celite. (Note: The filter cake must be kept wet with solvent because the palladium saturated with hydrogen is pyrophoric in air when dry.) The hexane was evaporated and the residue was purified by flash column chromatography (silica, EtOAc: Hexanes / 15:
85) affording 2.82 g (92 % yield) of (Z)-oct-2-en-1-ol 16 as a colorless oil. Identity and purity of the product were confirmed by 1H and 13C NMR. Spectroscopic data were found
31 1 to match lit. data. H NMR (400 MHz, CDCl3) δ 0.88 (t, 1H, J = 6.8 Hz), 1.23-1.41 (m,
6H), 2.07 (q, 2H, J = 7.2 Hz), 4.19 (d, 2H, J = 6.4 Hz), 5.47-5.68 (m, 2H); 13C NMR (100
MHz, CDCl3) δ 14.40, 22.86, 27.73, 29.62, 31.75, 58.96, 128.58, 133.67.
(Z)--(3-pentyloxiran-2-yl)methanol (17) (C8H16O2)
HO O
An oven-dried 250 mL, round-bottomed flask was equipped with a magnetic stirring bar. The flask was charged with m-chloroperoxybenzoic acid (m-CPBA) (7.9 g,
35.2 mmol, 77 % purity) and CH2Cl2 (80 mL). The reaction mixture was cooled to 0 °C
66
and (Z)-oct-2-en-1-ol (4.50 g, 35.2 mmol) in CH2C12 (5 mL) was added.The solution was warmed to room temperature and stirred for 2 h. The excess oxidant was quenched with a saturated solution of sodium thiosulfate (5.0 mL dropwise) and continued stirring for an additional 1 h. The reaction mixture was extracted with CH2C12 (3 × 30 mL), the
combined organic phases dried with anhydrous magnesium sulfate and concentrated
under reduced pressure. The crude product was distilled to give 4.82 g (95%) of (Z)-(3-
pentyloxiran-2-yl)methanol 17 (bp 60 oC, 0.2 Torr) as a colorless oil. Identity and purity of the product were confirmed by 1H and 13C NMR. Spectroscopic data were found to
32 1 match lit. data. H NMR (400 MHz, CDCl3) δ 0.90 (t, 3H, J = 7.2 Hz), 1.30-1.35 (m,
4H), 1.45-1.60 (m, 4H), 3.00-3.06 (m, 1H), 3.16 (dt, 1H, J = 7.2, 4.0 Hz), 3.68 (dd, 1H, J
13 = 7.2, 12.0 Hz), 3.86 (dd, 1H, J = 1.6, 12.0 Hz); C NMR (100 MHz, CDCl3) δ 14.32,
22.89, 26.66, 28.27, 31.92, 57.17, 57.70, 61.29.
(Z)-3-pentyloxirane-2-carbaldehyde (18) (C8H14O2)
O O
An oven-dried 250-mL, round-bottomed flask was equipped with a magnetic stirring bar and a rubber septum. The flask was charged with pyridinium chlorochromate
(12.26 g, 57 mmol) and 85 mL of dichloromethane. A solution of (3-pentyloxiran-2- yl)methanol 17 ( 4.10 g, 28.5 mmol) in 15 mL of dichloromethane was transferred into the reaction mixture via syringe over 5 min. The reaction mixture turned dark brown. The resulting mixture was stirred at ambient temperature for 3 h. The reaction mixture was then charged with 60 mL of diethyl ether and silica gel (30 g). The solution was decanted, and the remaining dark brown resinous polymer was thoroughly washed with three 30-
67
mL portions of diethyl ether. The combined dark brown / black ether was filtered and
concentrated by rotary evaporation at room temperature. Purification by column
chromatography (Hexanes: ether / 70: 30) afforded 2.23 g (55 % yield) of (Z)-3- pentyloxirane-2-carbaldehyde 18 as colorless oil. Identity and purity of the product were
1 13 1 confirmed by H and C NMR. H NMR (400 MHz, CDCl3) δ 0.89 (t, 3H, J = 7.2 Hz),
1.29-1.36 (m, 6H), 1.60-1.80 (m, 2H), 3.23-3.28 (m, 1H), 3.34 (t, 1H, J = 4.8 Hz), 9.46
13 (d, 1H, J = 5.2 Hz); C NMR (100 MHz, CDCl3) δ 14.24, 22.78 , 26.59, 28.42, 31.66,
58.26, 59.56, 199.55.
Methyl (E)-9-oxo-11-(3-pentyloxiran-2-yl)undec-10-enoate (19) (C19H32O4)
O O
O O
An oven-dried 50-mL, round-bottomed flask was equipped with a magnetic
stirring bar. The flask was charged with methyl 9-oxo-10-(triphenyl-l5-
phosphanylidene)decanoate 10 (1.11 g, 2.41 mmol) and CH2Cl2 (12 mL). The resulting
mixture was cooled to 0 °C in an ice bath. Then, cis-3-pentyloxirane-2-carbaldehyde 18
(291 mg, 2.41 mmol) was added in CH2Cl2 (12 mL) via syringe. The reaction mixture
continued to stir for an additional 4 h at 0 °C. Water (12 mL) was added to the mixture
and extracted with CH2Cl2 (3 × 25 mL).The combined organic phases were dried with
anhydrous magnesium sulfate and concentrated under reduced pressure. Purification by
column chromatography (Hexanes: ether / 85: 15) afforded 398 mg (51 %) of methyl (E)-
9-oxo-11-(3-pentyloxiran-2-yl)undec-10-enoate 19 as a yellow oil. Identity and purity of
the product were confirmed by 1H and 13C NMR. Spectroscopic data were found to match
68
16 1 lit. data. H NMR (500 MHz, CDCl3) δ 0.89 (t, 3H, J = 7.0 Hz), 1.25-1.68 (m, 18H),
2.34 (m, 2H), 2.53 (t, 1H, J = 7.0 Hz), 3.19 (m, 1H), 3.51 (dd, 1H, J = 6.0, 5.0 Hz), 3.65
(s, 1H), 6.41 (d, 1H, J = 16.0 Hz), 6.62 (dd, 1H, J = 16.0, 7.0 Hz); 13C NMR (125 MHz,
CDCl3) δ 14.26, 22.83, 24.30, 25.21, 26.30, 27.93, 29.26, 29.36, 29.38, 31.79, 34.64,
41.18, 55.81, 60.19, 133.18, 140.04, 174.15, 199.64.
(E)-9-oxo-11-(3-pentyloxiran-2-yl)undec-10-enoic acid( cis-EKODE-(E)-
Ib)(C18H30O4)
O O
HO O
Following the general procedure for PPL hydrolysis of methyl (E)-9-oxo-11-(3- pentyloxiran-2-yl)undec-10-enoate 19 afforded 29.0 mg (93 %) of cis-EKODE-(E)-Ib as a yellow solid. Identity and purity of the product were confirmed by 1H and 13C NMR.
16 1 Spectroscopic data were found to match lit. data. H NMR (500 MHz, CDCl3) δ 0.89 (t,
3H, J = 7.0 Hz), 1.25-1.68 (m, 18H), 2.34 (t, 2H, J = 7.5 Hz), 2.54(t, 1H, J = 7.5 Hz),
3.20 (m, 1H), 3.52 (ddd, 1H, J = 6.0, 4.5, 0.5 Hz), 6.40 (dd, 1H, J = 15.5, 0.5 Hz),
13 6.65(dd, 1H, J = 15.5, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 14.28, 22.85, 24.30,
24.93, 26.31, 27.94, 29.18, 29.32, 29.35, 31.81, 34.17, 41.17, 55.85, 60.24, 133.20,
140.09, 179.28, 199.71.
69
1,3-dibromopropan-2-one (20) (C3H4Br2O)
O Br Br
An oven-dried 2.0-L, round-bottomed flask was equipped with a mechanical stirrer. The flask was charged with 1,3-dichloro-2-propanone (25 g, 196 mmol) in
acetone (750 mL). Powdered lithium bromide (150 g, 1.73mol) was added slowly using a funnel. The reaction mixture was stirred at ambient temperature for 48 h. Additional
lithium bromide (100 g, 1.15 mol) and acetone (250 mL) were added and stirring
continued for an extra 24 h. Evaporation of the solvent gave a white solid that was
transferred to a 2.0 L separatory funnel charged with 1.0 L of cold water. The aqueous
phase was extracted with methylene chloride (3 × 500 mL), the combined organic phases
were dried with anhydrous magnesium sulfate and concentrated under reduced pressure
to give 42.1 g (99 % crude yield) of the 1,3-dibromopropan-2-one as a yellow syrup.
(Caution: The product has a low boiling point. The water bath in rota-vap needs to be
cold to avoid losing the product. bp79.5-805. oC at 9 Torr).33 Analysis of the crude by 1H
NMR indicates conversion was > 90 %. Identity and purity of the product were
confirmed by 1H NMR and 13CNMR. Spectroscopic data from 1HNMR were found to
33 1 13 match lit. data. H NMR (400 MHz, CDCl3) δ 4.13; C NMR (100 MHz, CDCl3) δ
31.18, 194.13.
70
1-bromo-3-(triphenylphosphoranylidene)propan-2-one (21) (C21H18BrOP)
O Ph P Br 3
An oven-dried 500-mL, two-necked round-bottomed flask containing a magnetic
stirring bar was equipped with an equalizing dropping funnel and an argon inlet. The flask was charged with triphenylphosphine (49.8 g, 190 mmol) in toluene (125 mL). The solution of crude 1,3-dibromopropan-2-one (41.0 g, 190 mmol) in toluene (125 mL) was added through the dropping funnel. Stirring continued overnight at ambient temperature.
The white precipitate formed during the reaction was collected by filtration, washed with toluene, and concentrated under reduced pressure. Powdered sodium bicarbonate (42 g,
0.5 mol) was added to a stirred solution of dried salt in 60% aqueous methanol (800 mL)
was added. Stirring continued for an additional 30 min and more water (200 mL) was
added to the mixture. After being stirred for another 30 min, the solid was collected by
filtration, and thoroughly washed with water. The solid was transferred to a 2.0 L
separatory funnel charged with 1.0 L of cold water. The aqueous phase was extracted with methylene chloride (3 × 500 mL), the combined organic phases were dried with
anhydrous sodium sulfate and concentrated under reduced pressure to give 62.6 g (83%
crude yield) of the 1-bromo-3-(triphenylphosphoranylidene)propan-2-one as a white solid. Analysis of the crude by 1H NMR indicates conversion was >60% (contaminated
with 1-chloro-3-(triphenylphosphoranylidene)propan-2-one). Identity and purity of the
1 13 1 product were confirmed by H and C NMR and HRMS. H NMR (400 MHz, CDCl3) δ
4 2 13 3.91 (d, 2H, JHP = 1.6), 4.26 (d, 1H, JHP = 24.0), 7.42-7.70 (15H,m); C NMR (100
3 1 1 MHz, CDCl3) δ 35.77 (d, JCP = 17.4 Hz), 52.67 (d, JCP = 109.0 Hz), 125.95 (d, JCP = 56
3 4 2 Hz, C-1’), 129.22 (d, JCP = 12.3 Hz, C-3’) 132.65 (d, JCP = 2.8 Hz, C-4’) 133.36 (d, JCP
71
2 = 10.2 Hz, C-2’), 184.99 (d, JCP = 4.3 Hz), HRMS (ESI) m/z calcd for C21H19BrOP+
(M+1)+ 397.03514, found 397.03520.
Dimethyl(2-oxo-3-(triphenylphosphoranylidene)propyl)sulfonium
tetrafluoroborate [Bifunctional conjunctive ylide] (22) (C23H24BF4OPS)
O Ph P - 3 S(CH3)2 BF4
An oven-dried 1.0L, round-bottomed flask was equipped with a magnetic stirring bar. The flask was charged with sodium tetrafluoroborate (27.5 g, 0.25 mol), methyl
sulfide (15.5 g, 0.25 mol), 1-bromo-3-(triphenylphosphoranylidene)propan-2-one (19.86 g, 50 mmol) and acetone (500 mL). The reaction mixture continued to stir at ambient temperature for 48h. The filtrate was concentrated under reduced pressure. Purification by column chromatography (DCM: MeOH / 95: 5) afforded 17.95 g (77 % yield) of bifunctional conjunctive ylide as a white solid. Identity and purity of the product were confirmed by 1H and 13C NMR and HRMS.13C NMR data did not duplicate lit. data.23 1H
2 NMR (400 MHz, CDCl3) δ 2.91 (s,6H), 4.16 (d, 1H, JHP = 21.2 Hz), 4.42 (s, 2H), 7.47-
13 3 7.64 (15H, m); C NMR (100 MHz, CDCl3) δ 25.01, 52.43 (d, JCP = 20.5 Hz), 56.71 (d,
1 1 3 JCP = 104 Hz), 125.20 (d, JCP = 90.8 Hz, C-1’), 129.45 (d, JCP = 12.4 Hz, C-3’), 133.09
2 2 (bs, C-4’), 133.21 (d, JCP = 10.3 Hz, C-2’), 177.06 (d, JCP = 4.0 Hz), HRMS (ESI) m/z
+ + calcd for C23H24OPS (M+1) 379.12800, found 379.12807.(confirms the cation)
72
Hex-2-yn-1-ol (28) (C6H10O)
HO
An oven-dried 500-mL, two-necked round-bottomed flask equipped with a
magnetic stirring bar was sealed under argon with two rubber septa, one of which contained a needle adapter attached to an argon-filled balloon. The flask was charged with propargyl alchohol (5.61 g, 0.1 mol), tetrahydrofurane (THF, 150 mL) and hexamethyl phosphoric triamide (HMPA, 40 mL). The solution was cooled in a dry ice- acetone bath at -78 °C, and n-butyllithium (84.4 mL, 2.37 M in hexane, 0.2mol) was added slowly via syringe. The resulting solution was stirred for 5 min at -78 °C, after which slowly warmed to -30 °C. Next, 1-bromopropane (9.23 g, 75 mmol) was added slowly to the mixture via syringe. The temperature was slowly increased to ambient temperature overnight while the mixture stirred. The reaction mixture was neutralized with aqueous saturated NH4Cl (100 mL) and extracted with EtOAc (3 × 100 mL), and the combined organic phases were dried with anhydrous sodium sulfate and concentrated under reduced pressure. Purification by column chromatography (EtOAc: Hexanes / 85:
15) afforded 6.50 g (57 %) of product as a colorless oil. Identity and purity of the product
were confirmed by 1H and 13C NMR. Spectroscopic data were found to match lit. data.34
1 H NMR (400 MHz, CDCl3) δ 0.96 (t, 1H, J = 7.2 Hz), 1.51 (sextet, 2H, J = 7.2 Hz), 1.99
(bs, 1H), 2.17 (tt, 2H, J = 7.2, 2.0 Hz), 4.24 (dt, 2H, J = 5.6, 2.0 Hz); 13C NMR (100
MHz, CDCl3) δ 13.72, 20.96, 22.35, 51.41, 78.75, 86.44.
73
Hex-5-yn-1-ol 29 (C6H10O)
HO
An oven-dried 500-mL, two-necked round-bottomed flask equipped with a
magnetic stirring bar was sealed under argon with two rubber septa, one of which contained a needle adapter attached to an argon inlet. The flask was charged with Li (4.48 g, 0.64mol) and 1,3- diaminopropane (240 mL). The mixture was stirred while heating in an oil bath at 70 °C until the blue color discharged (1 h). The prepared lithium amide
suspension was cooled to room temperature. Next, potassium tert-butoxide (42.4 g, 384 mmol), was added to the flask using a powder funnel. The resulting pale yellow solution was stirred for 20 min at room temperature, and then Hex-2-yn-1-ol (6.27 g, 64 mmol) was added over 10 min via syringe. The reddish-brown mixture was stirred for 3h and then poured into ice-water (1L) and extracted with Et2O (3 × 300 mL). The ether extracts
were combined and successively washed with 1 L of water, 10 % hydrochloric acid and
saturated sodium chloride solution. The ether solution was dried over anhydrous
magnesium sulfate, filtered and concentrated under reduced pressure. Purification by
column chromatography (EtOAc: Hexanes / 80: 20) afforded 5.33 g (84 %) of product as
a colorless oil. Identity and purity of the product were confirmed by 1H and 13C NMR.
35 1 Spectroscopic data were found to match lit. data. H NMR (400 MHz, CDCl3) δ 1.54-
1.68 (m, 4H), 1.93 (t, 1H, J = 2.8 Hz), 2.19 (td, 2H, J = 6.8, 2.8 Hz), 2.56 (bs, 1H), 3.60
13 (t, 2H, J = 6.4 Hz); C NMR (100 MHz, CDCl3) δ 18.40, 24.95, 31.82, 62.31, 68.82,
84.58.
74
Hex-5-ynal (30) ( C6H12O)
O
An Oven-dried 500-mL, three-necked, round-bottomed flask was equipped with a
magnetic stirring bar, a rubber septum, a glass stopper and an argon inlet. The flask was
charged with pyridinium chlorochromate (21.5 g, 100 mmol) and dichloromethane (150
mL). A solution of Hex-5-yn-1-ol (4.9 g, 50 mmol) in 20 mL of dichloromethane was
transferred into the reaction mixture via syringe over 5 min, and the resulting mixture
was stirred at ambient temperature for 3 h. The reaction mixture was then charged with
125 mL of diethyl ether and silica gel (50 g), the solution was decanted and the remaining
dark brown resinous polymer was thoroughly washed with three 50-mL portions of
diethyl ether. The combined dark brown / black ether was filtered and concentrated by
rotary evaporation at room temperature. The crude product was distilled to give 2.49 g
(52 %) of product (bp 29 oC, 1 Torr) as a colorless oil. Identity and purity of the product were confirmed by 1H and 13C NMR. Spectroscopic data were found to match lit. data.36
1 H NMR (400 MHz, CDCl3) δ 1.85 (quintet, 2H, J = 7.2 Hz), 1.97 (t, 1H, J = 2.8 Hz),
2.26 (td, 2H, J = 7.2, 2.8 Hz), 2.60 (td, 2H, J = 7.2, 1.2 Hz), 9.80 (1H, bs); 13C NMR (100
MHz, CDCl3) δ 18.07, 21.09, 42.82, 69.68, 83.48, 202.13.
Hexan-2,2,3,3,4,4,5,5,6,6,6-d11-1-ol (32) (C6H3D11O)
D D D D
HO CD3 D D D D
An oven-dried 250-mL, three-necked round-bottomed flask was equipped with a
glass stopper, a dropping funnel, a reflux condenser topped with an drying tube and a
75
magnetic stirring bar. The flask was charged with LiAlH4 (1.4 g, 37 mmol) and
anhydrous ether (50 mL). A solution of hexanoic-d11 acid (3.2 g, 25 mmol) in ether (50 mL) was transferred to the addition funnel and added over a 10-min period. The reaction mixture was heated under reflux for another 2h, diluted with ether (50 mL) and quenched by addition of water (25 mL) slowly via syringe. Next, the mixture was extracted with
EtOAc (3 × 50 mL). The combined organic phases were dried with anhydrous sodium sulfate and concentrated under reduced pressure affording 2.75 g (97 %) of product as a colorless oil. Identity and purity of the product were confirmed by 1H and 13C NMR. 1H
13 NMR (400 MHz, CDCl3) δ 1.81 (s, 1H, OH), 3.60 (s, 2H); C NMR (100 MHz, CDCl3)
δ 62.90.
Hexanal-2,2,3,3,4,4,5,5,6,6,6-d11 (33) (C6HD11O)
D D D D
O CD3 D D D D
An oven-dried 250-mL, three-necked round-bottomed flask was equipped with a magnetic stirring bar, a rubber septum, a glass stopper and an argon inlet. The flask was charged with pyridinium chlorochromate (8.6 g, 40 mmol) and dichloromethane (60 mL).
A solution of hexan-2,2,3,3,4,4,5,5,6,6,6-d11-1-ol ( 2.26 g, 20 mmol) in dichloromethane
(10mL) was transferred into the reaction mixture via syringe over 5 min, and the resulting
mixture was stirred at ambient temperature for 3 h. The reaction mixture was then charged with diethyl ether (50 mL) and silica gel (20 g). The solution was decanted, and the remaining dark brown resinous polymer thoroughly washed with three 20-mL portions of diethyl ether. The combined dark brown / black ether was filtered and
76
concentrated by rotary evaporation at room temperature to give 1.51 g (68 %) of the
crude product as a yellow oil. The product was used with no further purification. Identity
and purity of the product were confirmed by 1H and 13C NMR. 1H NMR (400 MHz,
13 CDCl3) δ 9.74 (1H, CHO); C NMR (100 MHz, CDCl3) δ 203.66.
General procedure: Johnson–Corey–Chaykovsky reaction
An oven-dried 50-mL, two-necked round-bottomed flask containing a magnetic stirring bar was sealed under argon with two rubber septa, one of which contained a needle adapter attached to an argon-inlet. The solution was cooled in a dry ice-ethanol bath at -30 °C. The flask was charged with bifunctional conjunctive ylide (2.0 mmol) and sodium hydride (80 mg, 2.0 mmol, 60 % in mineral oil) and desired aldehyde (2.0 mmol) and acetonitrile (30 mL). The reaction mixture continued to stir while bring heat to room temperature over 3 h. The reaction mixture was neutralized with aqueous saturated
NH4Cl (20 mL) and extracted with EtOAc (3 × 20 mL), the combined organic phases
were dried with anhydrous sodium sulfate and concentrated under reduced pressure.
Purification by column chromatography (DCM: Methanol / 98: 2) afforded desired epoxy
products.
77
Methyl 8-(3-(2-(triphenyl- λ 5-phosphanylidene)acetyl)oxiran-2-yl)octanoate
(23) (C31H35O4P)
O O Ph3P O O
Following the general procedure for Johnson–Corey–Chaykovsky reaction and
using aldehyde 2 (372 mg, 2.0 mmol) and completion in 8 h. Purification by column
chromatography afforded 583 mg (58%) of product as a yellow oil. Identity and purity of
1 13 1 the product were confirmed by H, C NMR and HRMS. H NMR (400 MHz, CDCl3) δ
1.20-1.79 (m, 12H), 2.28 (t, 2H, J = 7.6 Hz), 3.05 (bm, 1H), 3.14 (t, 1H, J = 2 Hz), 3.65
2 13 (s, 3H), 3.94 (d, 1H, JHP = 25.6 Hz), 7.41-7.71 (m, 15H); C NMR (100MHz, CDCl3) δ
1 25.03, 26.11, 29.18, 29.26, 29.30, 32.23, 34.19, 48.26 (d, 1H, JCP = 98 Hz), 51.58, 59.90,
1 3 60.04 (d, J = 17 Hz), 126.56 (d, JCP = 91 Hz, C-1’), 129.05 (d, JCP = 12.3 Hz, C-3’),
4 2 2 132.40 (d, JCP = 2.7 Hz, C-4’), 133.17 (d, JCP = 10.3 Hz, C-2’), 174.40, 188.11 (d, JCP =
+ + 4.0 Hz), HRMS (ESI) m/z calcd for C31H36O4P (M+1) 503.23457, found 503.23469.
1-(3-pentyloxiran-2-yl)-2-(triphenyl-λ5-phosphanylidene)ethan-1-one (26)
(C27H29O2P)
O Ph3P O
Following the general procedure for Johnson–Corey–Chaykovsky reaction and
using aldehyde 24 (200 mg, 2.0 mmol) and purification by column chromatography
afforded 600 mg (72 %) of product as a yellow oil. Identity and purity of the product
1 13 1 were confirmed by H, C NMR and HRMS. H NMR (500 MHz, CDCl3) δ 0.88 (t, 3H,
78
2 J = 6.5 Hz), 1.22-1.82 (m, 8H), 3.05 (bm, 1H), 3.13 (t, 1H, J = 2.0 Hz), 3.94 (d, 1H, JHP=
13 24 Hz), 7.35-7.73 (m, 15H); C NMR (125 MHz, CDCl3) δ14.24, 22.80, 25.95, 31.82,
1 1 32.35, 48.86 (d, JCP = 110 Hz), 60.07, 60.17 (d, J = 16.6 Hz), 126.36 (d, JCP = 90.6 Hz,
3 4 2 C-1’), 129.15 (d, JCP = 12.3 Hz, C-3’), 132.49 (d, JCP = 2.8 Hz, C-4’), 133.31 (d, JCP =
2 + + 10.2 Hz, C-2’), 188.31 (d, JCP = 3.5 Hz), HRMS (ESI) m/z calcd for C27H30O2P (M+1)
417.19779, found 417.19787.
1-(3-(pent-4-yn-1-yl)oxiran-2-yl)-2-(triphenyl- λ5-phosphanylidene)ethan-1-
one (34) (C27H25O2P)
O Ph3P O
Following the general procedure for Johnson–Corey–Chaykovsky reaction and
using aldehyde 30 (192 mg, 2.0 mmol) and purification by column chromatography
afforded 553 mg (67 %) of product as a yellow oil. Identity and purity of the product
1 13 1 were confirmed by H, C NMR and HRMS. H NMR (500 MHz, CDCl3) δ 1.49-1.71
(m, 4H), 1.89 (t, 1H, J = 2.0 Hz), 2.16-2.24 (m, 2H), 3.05 (ddd, 1H, J = 8.5, 4, 2.0 Hz),
2 13 3.13 (t, 2H, J = 2.0 Hz), 3.99 (d, 1H, JHP = 23.5 Hz), 7.34-7.63 (m, 15H); C NMR (125
1 MHz, CDCl3) δ 18.25, 25.06, 31.06, 49.07 (d, 1H, JCP = 110 Hz), 59.10, 59.72 (d, J = 17
1 3 Hz), 68.70, 84.03, 126.44 (d, JCP = 91 Hz, C-1’), 129.01 (d, JCP = 12.4 Hz, C-3’), 132.38
4 2 2 (d, JCP = 2.8 Hz, C-4’), 133.15 (d, JCP = 10.3 Hz, C-2’), 187.65 (d, JCP = 4.0 Hz).
79
1-(3-(pentyl-d11)oxiran-2-yl)-2-(triphenyl-l5-phosphanylidene)ethan-1-one
(35) (C27H18D11O2P)
O D D D D Ph3P CD O 3 D D D D
Following the general procedure for Johnson–Corey–Chaykovsky reaction and
using aldehyde 33 (223 mg, 2.0 mmol) and purification by column chromatography
afforded 599 mg (70 %) of product as a brown oil. Identity and purity of the product were
1 13 1 confirmed by H, C NMR and HRMS. H NMR ( 500 MHz, CDCl3) δ 3.04 (b, 1H),
2 13 3.04 (t, 1H, J = 2.0 Hz), 3.94 (d, 1H, JHP = 24.0 Hz), 7.39-7.72 (m, 15 H); C NMR (125
1 1 MHz, CDCl3) δ 48.90 (d, JCP = 110.0 Hz), 60.09, 60.13 (d, J = 16.5 Hz), 126.79 (d, JCP
3 4 = 90.6 Hz, C-1’), 129.48 (d, JCP = 12.3 Hz, C-3’) 132.37 (d, JCP = 2.8 Hz, C-4’) 133.36
2 2 (d, JCP = 10.3 Hz, C-2’), 188.44 (d, JCP = 4.0 Hz), HRMS (ESI) m/z calcd for
+ + C27H19D11O2P (M+1) 428.267, found 428.266.
General procedure: Wittig reaction
An oven-dried 50-mL, round-bottomed flask, containing a magnetic stirring bar
charged with desired epoxy (1 mmol) from Johnson–Corey–Chaykovsky reaction and
desired aldehyde (1 mmol) and toluene (10 mL). The mixture was heated under reflux overnight and allowed to cool to room temperature. The reaction mixture was concentrated under reduced pressure. Purification by column chromatography (Hexane:
Et2O / 85: 15) afforded α, β-unsaturated keto epoxide products.
80
Methyl (E)-8-(3-(oct-2-enoyl)oxiran-2-yl)octanoate (25) (C19H32O4)
O O
O O
Following the general procedure for Wittig reaction between 23 and 24 and
purification by column chromatography afforded 278 mg (86%) of product as a yellow
oil. Identity and purity of the product were confirmed by 1H, 13C NMR and HMRS.
16 1 Spectroscopic data were found to match lit. data. H NMR (400 MHz, CDCl3) δ 0.87 (t,
3H, J = 7.2 Hz), 1.25-1.69 (m, 18H), 2.18 (dd, 2H, J = 15.0, 7.0 Hz), 2.30 (t, 2H, J = 7.5
Hz), 3.02(t, 1H, J = 6 Hz), 3.32 (bs, 1H), 3.67(s, 3H), 6.23 (d, 1H, J = 15.5 Hz), 7.08
13 (1H, dt, J = 15.5, 7.0 Hz); C NMR (100 MHz, CDCl3) δ 14.32, 22.76, 25.11, , 26.07,
27.97, 29.31, 29.41, 29.42, 31.71, 32.14, 33.08, 34.38, 51.83, 58.66, 59.36, 124.27,
+ + 151.02, 174.61. 196.05; HRMS (ESI) m/z calcd for C19H32O4Na (M+Na) 347.21928,
found 347.21943.
(E)-8-(3-(oct-2-enoyl)oxiran-2-yl)octanoic acid (trans-EKODE-(E)-
IIa)(C18H30O4)
O O
OH O
Following the general procedure for PPL hydrolysis of methyl (E)-8-(3-(oct-2-
enoyl)oxiran-2-yl)octanoate 25 afforded 146 mg (94 %) of product as a yellow oil.
Identity and purity of the product were confirmed by 1H, 13C NMR and HRMS.
16 1 Spectroscopic data were found to match lit. data. H NMR (500 MHz, CDCl3) δ 0.89 (t,
3H, J = 7.0 Hz), 1.21-1.74 (m, 18H), 2.17-2.24 (m, 2H), 2.34 (t, 2H, J = 7.0 Hz), 3.04
81
(ddd, 1H, J = 6.0, 5.0, 2.0 Hz), 3.33 (d, 1H, J = 2.0 Hz), 6.23 (dt, 1H, J = 15.5, 1.5 Hz),
13 7.07 (1H, dt, J = 15.5, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 14.27, 22.73, 24.91,
26.08, 27.94, 29.20, 29.36, 29.38, 31.68, 32.11, 33.06, 34.25, 58.66, 59.33, 124.26,
- - 151.06, 179.81. 196.08; HRMS (FAB) calcd for C18H29O4 (M-H) 309.427, found
309.207.
Methyl (E)-8-(3-(oct-2-en-7-ynoyl)oxiran-2-yl)octanoate (36) (C19H28O4)
O O
O O
Following the general procedure for Wittig reaction between 23 and 30 and
purification by column chromatography afforded 260 mg (81 %) of product as a yellow oil. Identity and purity of the product were confirmed by 1H, 13C NMR and HRMS. 1H
NMR (500 MHz, CDCl3) δ 1.26-1.66 (m, 12H), 1.70 (quintet, 2H, J = 7.0 Hz), 1.98 (t,
1H, J = 2.5 Hz), 2.22 (td, 2H, J = 7.0, 2.5 Hz), 2.30 (t, 2H, J = 7.0 Hz), 2.36 (t, 2H, J =
7.0 Hz), 3.04 (td, 1H, J = 5.5, 2.0 Hz), 3.33 (d, 1H, J = 2 Hz), 3.66 (s, 3H), 6.27 (d, 1H, J
13 = 16 Hz), 7.06 (1H, dt, J = 16.0, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 18.27, 25.20,
26.09, 29.30 (2C), 29.39, 29.41, 31.81, 32.12, 34.38, 51.82, 58.61, 59.42, 69.49, 83.716,
+ + 124.76, 149.27, 174.59, 195.93; HRMS (ESI) m/z calcd for C19H28O4Na (M+Na)
343.18798, found 343.18812.
(E)-8-(3-(oct-2-en-7-ynoyl)oxiran-2-yl)octanoic acid (40) (C18H26O4)
O O
OH O
82
Following the general procedure for PPL hydrolysis of methyl (E)-8-(3-(oct-2-en-
7-ynoyl)oxiran-2-yl)octanoate 36 afforded 139 mg (91 %) of product as yellow oil.
Identity and purity of the product were confirmed by 1H, 13C NMR and HRMS. 1H NMR
(500 MHz, CDCl3) δ 1.22-1.88 (m, 12H), 1.70 (quintet, 2H, J = 7.0 Hz), 1.98 (t, 1H, J =
2.5 Hz), 2.17-2.29 (m, 4H), 2.30-240 (m, 2H), 3.05 (t, 1H, J = 5.0 Hz), 3.33 (bs, 1H),
13 6.27 (d, 1H, J = 15.5 Hz), 7.06 (1H, dt, J = 15.5, 7.0 Hz); C NMR (125 MHz, CDCl3) δ
18.27, 24.92, 26.08, 26.96, 29.20, 29.36, 29.38, 31.82, 32.09, 34.15, 58.62, 59.41, 69.50,
- - 83.72, 124.75, 149.33, 179.28, 195.97; HRMS (ESI) m/z calcd for C18H25O4 (M-H)
305.395, found 305.176.
Methyl (E)-8-(3-(oct-2-enoyl-4,4,5,5,6,6,7,7,8,8,8-d11)oxiran-2-yl)octanoate
(37) (C19H21D11O4)
D D D D O O D C O 3 O D D D D
Following the general procedure for Wittig reaction between 23 and 33 and
purification by column chromatography afforded 245 mg (73 %) of product as a yellow oil. Identity and purity of the product were confirmed by 1H and 13C NMR and HMRS.
1 H NMR (500 MHz, CDCl3) δ 1.27-1.71 (m, 12H), 2.30 (t, 2H, J = 7.5 Hz), 3.04 (t, 1H, J
= 5.0 Hz), 3.33 (bs, 1H), 3.66 (s, 3H), 6.23 (d, 1H, J = 16.0 Hz), 7.07 (d, 1H, J = 16.0
13 Hz); C NMR (125 MHz, CDCl3) δ 25.20, 26.09, 29.30, 29.39, 29.41, 32.13, 34.37,
51.81, 58.64, 59.34, 124.34, 150.95, 174.59, 196.02; HRMS (ESI) m/z calcd for
+ + C19H25D11NO4 (M+NH4) 353.567, found 353.333.
83
(E)-8-(3-(oct-2-enoyl-4,4,5,5,6,6,7,7,8,8,8-d11)oxiran-2-yl)octanoic acid (41)
(C18H19D11O4)
D D D D O O D C OH 3 O D D D D
Following the general procedure for PPL hydrolysis of methyl (E)-8-(3-(oct-2-
enoyl-4,4,5,5,6,6,7,7,8,8,8-d11)oxiran-2-yl)octanoate 37 afforded 150 mg (93 %) of product as yellow oil. Identity and purity of the product were confirmed by 1H, 13C NMR
1 and HMRS. H NMR (500 MHz, CDCl3) δ 1.29-1.71 (m, 12H), 2.28 (t, 2H, J = 7.5 Hz),
3.01 (m, 1H), 3.52 (bs, 1H), 6.28 (d, 1H, J = 16 Hz), 7.10 (d, 1H, J = 16.0 Hz); 13C NMR
(125 MHz, CDCl3) δ 26.82, 26.03, 30.10, 30.23, 30.24, 32.82, 34.91, 59.43, 60.01,
- - 126.55, 152.05, 177.68, 197.45; HRMS (ESI) m/z calcd for C18H18D11O4 (M-H)
320.493, found 320.276.
Methyl (E)-11-oxo-11-(3-pentyloxiran-2-yl)undec-9-enoate (27) (C19H32O4)
O O
O O
Following the general procedure for Wittig reaction between 26 and 24 and
purification by column chromatography afforded 234 mg (72 %) of product as a yellow
oil. Identity and purity of the product were confirmed by 1H, 13C NMR and HMRS.
16 1 Spectroscopic data were found to match lit. data. H NMR (500 MHz, CDCl3) δ 0.87 (t,
3H, J = 7.2 Hz), 1.19-1.73 (m, 18H), 2.21 (dd, 2H, J = 7.0, 2 Hz), 2.30 (t, 2H, J = 7.5
Hz), 3.05 (td, 1H, J = 5.5, 2.0 Hz), 3.34 (d, 1H, J = 2.0 Hz), 3.67 (s, 3H), 6.23 (d, 1H, J =
13 15.5 Hz), 7.07 (1H, dt, J = 15.5, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 14.27, 22.83,
84
25.19, 25.83, 28.19, 29.22, 29.31 (2C), 31.76, 32.13, 33.04, 34.37, 51.83, 58.75, 59.38,
+ + 124.32, 150.82, 174.64, 196.11; HRMS (ESI) m/z calcd for C19H32O4Na (M+Na)
347.21928, found 347.21942.
(E)-11-oxo-11-(3-pentyloxiran-2-yl)undec-9-enoic acid(trans-EKODE-(E)-
IIb) (C18H28O4)
O O
OH O
Following the general procedure for PPL hydrolysis of methyl (E)-11-oxo-11-(3- pentyloxiran-2-yl)undec-9-enoate 27 afforded 143 mg (92 %) of product as yellow oil.
Identity and purity of the product were confirmed by 1H, 13C NMR and HRMS.
16 1 Spectroscopic data were found to match lit. data. H NMR (400 MHz, CDCl3) δ 0.88 (t,
3H, J = 7.2 Hz), 1.17-1.67 (m, 18H), 2.21 (dq, 2H, J = 6.8, 1.6 Hz), 2.33 (t, 2H, J = 7.6
Hz), 3.05 (ddd, 1H, J = 6.0, 5.0, 2.0 Hz), 3.34 (d, 1H, J = 2.0 Hz), 6.25 (dt, 1H, J = 15.5,
13 1.5 Hz), 7.06 (1H, dt, J = 15.5, 7.0 Hz); C NMR (100 MHz, CDCl3) δ 14.28, 22.84,
24.93, 25.83, 28.20, 29.22, 29.30 (2C), 31.77, 32.14, 33.04, 34.20, 58.74, 59.40, 124.32,
- - 150.76, 179.44, 196.09, HRMS (FAB) calcd for C18H29O4 (M-H) 309.427, found
309.207.
85
Methyl (E)-11-oxo-11-(3-(pent-4-yn-1-yl)oxiran-2-yl)undec-9-enoate (38) (
C19 H28O4)
O O
O O
Following the general procedure for Wittig reaction between 34 and 2 and
purification by column chromatography afforded 208 mg (65 %) of product as a yellow oil. Identity and purity of the product were confirmed by 1H and 13C NMR and HRMS.
1 H NMR (500 MHz, CDCl3) δ 1.24-1.90 (m, 16H), 1.97 (t, 1H, J = 2.5 Hz), 2.18-2.25 (m,
2H), 2.30 (t, 2H, J = 7.5 Hz), 3.07 (dt, 1H, J = 5.0, 1.5 Hz), 3.38 (d, 1H, J = 1.5 Hz), 3.66
(s, 3H), 6.23 (d, 1H, J = 16 Hz), 7.08 (1H, dt, J = 16.0, 7.0 Hz); 13C NMR (125 MHz,
CDCl3) δ 18.39, 24.99, 25.20, 28.19, 29.22, 29.31, 30.97, 33.06, 34.36, 51.80, 58.08,
59.06, 69.47, 83.72, 124.41, 151.03, 174.53, 195.76; HRMS (ESI) m/z calcd for
+ + C19H28O4Na (M+Na) 343.18798, found 343.18812.
(E)-11-oxo-11-(3-(pent-4-yn-1-yl)oxiran-2-yl)undec-9-enoic acid (42) ( C18
H26O4)
O O
OH O
Following the general procedure for PPL hydrolysis of methyl (E)-11-oxo-11-(3-
(pent-4-yn-1-yl)oxiran-2-yl)undec-9-enoate 38 afforded 144 mg (94 %) of product as a yellow oil. Identity and purity of the product were confirmed by 1H, 13C NMR and
1 HRMS. H NMR (500 MHz, CDCl3) δ 1.22-1.88 (m, 12H), 1.97 (t, 1H, J = 2.5 Hz), 2.17-
2.32 (m, 4H,), 2.35 (t, 2H, J = 7.5 Hz), 3.08 (bs, 1H), 3.29(d, 1H, J = 5 Hz), 6.26 (d, 1H,
86
13 J = 15.5 Hz), 7.08 (1H, dt, J = 15.5, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 18.39,
24.93, 24.99, 28.18, 29.21, 29.28, 29.35, 30.97, 33.06, 34.09, 58.11, 59.11, 69.49, 83.74,
- - 124.43, 151.00, 178.90, 195.76; HRMS (ESI) m/z calcd for C18H25O4 (M-H) 305.395,
found 305.176.
Methyl (E)-11-oxo-11-(3-(pentyl-d11)oxiran-2-yl)undec-9-enoate (39) (C19
H21D11O4)
D D D D O O D C O 3 O D D D D
Following the general procedure for Wittig reaction between 35 and 2 and
purification by column chromatography afforded 146 mg (62 %) of product as yellow oil.
Identity and purity of the product were confirmed by 1H and 13C NMR and HRMS. 1H
NMR (500 MHz, CDCl3) δ 1.23-1.67 (m, 10H), 2.20 (q, 2H, J = 7.0 Hz), 2.27 (t, 2H, J =
7.0 Hz), 3.01 (s, 1H), 3.31 (s, 1H), 3.64 (s, 3H), 6.21 (d, 1H, J = 15.5 Hz), 7.05 (dt, 1H, J
13 = 15.5, 7.0 Hz); C NMR (125 MHz, CDCl3) δ 25.15, 28.15, 29.26, 29.27 (2C), 32.89,
34.31, 51.75, 58.58, 59.30, 124.28, 150.66, 174.51, 196.00 ,HRMS (ESI) m/z calcd for
+ + C19H25D11NO4 (M+NH4) 353.567, found 353.333.
(E)-11-oxo-11-(3-(pentyl-d11)oxiran-2-yl)undec-9-enoic acid (43)
D D D D O O D C OH 3 O D D D D Following the general procedure for PPL hydrolysis of methyl (E)-11-oxo-11-(3-
(pentyl-d11)oxiran-2-yl)undec-9-enoate 39 afforded 144 mg (94 %) of product as a
87
yellow oil. Identity and purity of the product were confirmed by 1H and 13C NMR and
1 HRMS. H NMR (500 MHz, CDCl3) δ 1.20-1.71 (m, 10H), 2.22 (q, 2H, J = 6.5 Hz), 2.35
(t, 2H, J = 6.5 Hz), 3.04 (s, 1H), 3.34 (s, 1H), 6.23 (d, 1H, J = 15.5 Hz), 7.07 (dt, 1H, J =
13 15.5, 6.0 Hz); C NMR (125 MHz, CDCl3) δ 24.95, 28.20, 29.23, 29.30, 29.36, 33.04,
34.17, 58.67, 59.36, 124.40, 150.79, 179.08, 196.16 HRMS (ESI) m/z calcd for
- - C18H18D11O4 (M-H) 320.493, found 320.276.
88
2. 6 References
(1) Samuelsson, B. The Journal of biological chemistry 2012, 287, 10070.
(2) Abrahamsson, S.; Bergstrom, S.; Samuelsson, B. P Chem Soc London 1962, 332.
(3) Austin, S. C.; Funk, C. D. Prostag Oth Lipid M 1999, 58, 231.
(4) Roberts, L. J.; Salomon, R. G.; Morrow, J. D.; Brame, C. J. Faseb J 1999, 13, 1157.
(5) Jacobs, A. T.; Marnett, L. J. Accounts Chem Res 2010, 43, 673.
(6) West, J. D.; Marnett, L. J. Chem Res Toxicol 2005, 18, 1642.
(7) Vila, A.; Tallman, K. A.; Jacobs, A. T.; Liebler, D. C.; Porter, N. A.; Marnett, L. J.
Chem Res Toxicol 2008, 21, 432.
(8) Gatbonton-Schwager, T. N.; Sadhukhan, S.; Zhang, G. F.; Letterio, J. J.; Tochtrop,
G. P. Redox Biol 2014, 2, 755.
(9) Yin, H.; Xu, L.; Porter, N. A. Chemical reviews 2011, 111, 5944.
(10) Zhu, X.; Tang, X.; Anderson, V. E.; Sayre, L. M. Chem Res Toxicol 2009, 22, 1386.
(11) Lundstrom, S. L.; Levanen, B.; Nording, M.; Klepczynska-Nystrom, A.; Skold, M.;
Haeggstrom, J. Z.; Grunewald, J.; Svartengren, M.; Hammock, B. D.; Larsson, B. M.; Eklund, A.;
Wheelock, A. M.; Wheelock, C. E. PloS one 2011, 6, e23864.
(12) Bruder, E. D.; Ball, D. L.; Goodfriend, T. L.; Raff, H. American journal of physiology. Regulatory, integrative and comparative physiology 2003, 284, R1631.
(13) Goodfriend, T. L.; Ball, D. L.; Egan, B. M.; Campbell, W. B.; Nithipatikom, K.
Hypertension 2004, 43, 358.
(14) Payet, M. D.; Goodfriend, T. L.; Bilodeau, L.; Mackendale, C.; Chouinard, L.;
Gallo-Payet, N. American journal of physiology. Endocrinology and metabolism 2006, 291,
E1160.
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(15) Wang, R.; Kern, J. T.; Goodfriend, T. L.; Ball, D. L.; Luesch, H. Prostaglandins,
leukotrienes, and essential fatty acids 2009, 81, 53.
(16) Lin, D.; Zhang, J.; Sayre, L. M. The Journal of organic chemistry 2007, 72, 9471.
(17) Curran, D. P. J Am Chem Soc 1983, 105, 5826.
(18) Kim, K. S.; Szarek, W. A. Can J Chem 1981, 59, 878.
(19) Minami, T.; Harui, N.; Taniguchi, Y. Journal of Organic Chemistry 1986, 51, 3572.
(20) Hercouet, A.; Lecorre, M. Tetrahedron 1977, 33, 33.
(21) Cichowicz, N. R.; Nagorny, P. Organic letters 2012, 14, 1058.
(22) Appel, R.; Mayr, H. Chemistry 2010, 16, 8610.
(23) Magdesieva, N. N.; Chovnikova, N. G.; Brunovlenskaya, I. I. Zh Org Khim+ 1984,
20, 2097.
(24) Magdesieva, N. N.; Kyandzhetsian, R. A.; Chovnikova, N. G.; Emelyanova, N. N.
Zh Org Khim+ 1981, 17, 340.
(25) Acharya, H. P.; Kobayashi, Y. Angewandte Chemie 2005, 44, 3481.
(26) Sun, M. J.; Deng, Y. J.; Batyreva, E.; Sha, W.; Salomon, R. G. Journal of Organic
Chemistry 2002, 67, 3575.
(27) Bally, I.; Gard, E.; Ciornei, E.; Biltz, M.; Balaban, A. T. J Labelled Compd 1975, 11,
63.
(28) Kusukawa, T.; Tanaka, S.; Inoue, K. Tetrahedron 2014, 70, 4049.
(29) Huisgen, R.; Rietz, U. Tetrahedron 1958, 2, 271.
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91
Chapter 3. Biological evaluation of EKODEs as potential activators of the
PPAR family of nuclear receptors
3.1 Peroxisome proliferator-activated receptors (PPARs) structure and
activity
Increasing dietary fat and sugar consumption in the Western diet along with the
sedentary lifestyle has expanded a number of pathological states such as diabetes, obesity and cardiovascular diseases (CVD).1 In fact, high caloric intake plays an important role in
limiting human lifespan through modulation of lipid metabolism, insulin sensitivity,
glucose homeostasis, etc.2 It is well-known that most species of higher multicellular
organisms such as humans have developed homeostatic mechanisms to be able to control
fatty acids intake and metabolism. A family of nuclear receptors, the peroxisome
proliferator-activated receptors (PPARs), makes a prominent contribution to regulate
lipid metabolism, catabolism and storage. In humans, the PPARs are a family of three receptors PPARα (NR1C1) (nuclear receptor subfamily 1, group C, member 1), PPARβ/δ
(NR1C2) and PPARγ (NR1C3) which are products of three different genes.3
Comparing to other types of nuclear receptors, PPARs share a domain structural feature. The most conserved regions are C and E in the protein sequence. The sequence homology in C shows the DNA binding domain (DBD) is highly conserved. However, in
region E, which represents the ligand binding-domains (LBD), moderate level of
conservation has been observed between subtypes. Interestingly, the differences in this
region have been contributed to distinct pharmacological effects of each subtype of
PPAR. The N-terminal A/B domain holds the AF1 ligand-independent activation domain,
and an AF2 ligand-dependent activation domain is situated in the F region. The numbers
92
shown in the LBD and DBD are the percentage amino-acid identity of human PPARβ/δ and PPARγ1 compared to human PPARα.4 (Figure 3.1)
A/B C D E F
Human PPARα N AF1 DBD LBD AF2 C
Human PPARβ/δ N 86 70 C
Human PPARγ N 83 68 C
Figure 3.1 PPARs motifs, conserved region in yellow, variable regions in red
The PPARs form a heterodimer with the retinoid X receptorα (RXRα) (NR2B)
and binds to peroxisome proliferator response elements (PPRE). These elements are
composed of two hexameric nucleotide recognition motifs of AGGTCA spaced by one
nucleotide. They reside on the promoter region of PPAR target genes. In addition, the
PPRE element consists of an additional AAACT motif at the upstream region. The
variable region D, or hinge region of PPAR, provides a broad interaction space with the
upstream AAACT element.5
In the absence of the ligand, the PPAR-RXR heterodimer remains bound to the
nuclear receptor corepressor (NCoR) and the silencing mediator of retinoid and thyroid
hormone receptor (SMRT), which are mostly present in the corepressor complex. SMRT
functions as a platform protein and facilitates the recruitment of histone deacetylases
(HDACs). In fact, the corepressor protein complex reduces target gene transcription by
causing the deacetylation of histones. Unlike the PPARβ/δ isoform, the PPARα and
PPARγ isoforms are not capable of binding to DNA while they are associated with the co-repressor complex. Similar to other types of nuclear receptors, the activity of these receptors can be enhanced by ligands; this process called “transactivation.”
93
Upon the binding of ligands to PPARs, they detach from the co-repressor complex. Next, the PPARs recruit specific co-activator complexes, such as steroid receptor co-activator 1 (SRC1) and cAMP response element binding (CREB)-binding protein (CBP)/p300. These co-activator complexes, through their histone- acetyltransferase activity, are capable of reorganizing the chromatin packaging and driving the transcriptional machinery to the promoter region. (Figure 3.2)
CBP/p300 CBP/p300 SRC1 SRC1
PPAR RXR 9cRA PPAR RXR 9cRA PPAR RXR 9cRA
NCoR/SMRT
Ligand PPRE PPRE PPRE
Figure3.2 PPAR binding to ligand along with RXR binding to 9-cis retinoic acid (9cRA) forms a complex which provides an on- off-switch for gene expression
The expression of PPARs is tissue-specific and regulated at different levels, including post-translational modification (PTM). In addition, their gene expression activity depends on the response elements’ position to which they bind. Furthermore, the availability of active chromatin can determine the level of target gene transcription.
Various types of fatty acids and related oxidized forms (eicosanoids) can bind to PPARs with variable degrees of specifity between isoforms.6
94
1 2 3 PPAR RXR 9cRA Co-activators Co-activators
PPAR RXR 9cRA PPAR RXR 9cRA
transcription factor
MAPK
Co-activators
transcription transcription transcription factor factor factor
Figure 3.3 Three main mechanisms proposed for negative regulation of transcription factors by
PPARs
Regulation of gene transcription with PPARs extends far beyond their effect on
transactivation of specific genes in an agonist-dependent manner. PPARs like other types
of nuclear receptors are also known to interact with various expression machineries and
act through transrepression. In fact, PPARs are capable of suppressing several important
signaling cascades. There are three main mechanisms: (1). PPARs compete on the common coactivators binding and consuming their limited supply for other genes, (2). they can bind to the transcription factors (for example, AP1, NF-κB, NFAT or STATs)
and make them unavailable for certain response elements, (3). they can inhibit mitogen-
activated protein kinase (MAPK) phosphorylation ability and inhibit transcription factor activity. For instance, PPARs can inhibit nuclear factor-κB (NFκB) and AP1 in
macrophages and deteriorate the pro-inflammatory signaling by decreasing the
expression of pro-inflammatory cytokines, chemokines and cell adhesion molecules.
(Figure 3.3)
95
3.2. The physiological functions of the PPARs
3.2.1 Functional role of PPARα
PPARα was the first member of the family discovered. Since it has a diverse
tissue distribution, it exhibits different physiological activities among organisms. In
humans, it mainly exists in metabolically active organs such as the liver, heart and
kidney. PPARα shows a central role in fatty acid catabolism and ketone body synthesis in the liver. Its expression is controlled by other transcription factors and nuclear receptors such as hepatocyte nuclear factor 4 (HNF4) and chicken ovalbumin upstream promoter- transcription factor II (COUP-TFII).
3.2.1.1 Exogenous ligands (synthetic Xenobiotics) of PPARα
Synthetic ligands such as fibrates (clofibrate, fenofibrate, GW7647) and Wy-
14,643 (pirinixic acid) are being used in the treatment of dyslipidemias. In addition, industrial chemicals and environmental pollutants exert their toxic effect through this receptor. For example, the plasticizers, such as di-(2-ethylhexyl)-phthalate (DEHP) and its related metabolite monoethylhexyl phthalate (MEHP), and di-(2-ethylhexyl) adipate
(DEHA) are activators of PPARα.7-9 Furthermore, certain perfluorinated compounds such
as perfluorooctanoic acid (PFOA) and perfluorooctanesulfonic acid (PFOS) also function
as PPARα ligands.10,11 (Figure 3.4)
96
O O S O O OH O OH N N O H O Cl Cl O Clofibrate Fenofibrate GW7647
O H O O N N S OH O O N OH Cl O O O Wy-14,643 DEHA MEHP O O F F F F F F F F F F F F F F O HO CCl3 HO CF S CF3 3 HO F F F F F F Trichloroacetic acid F F F F F F O PFOA PFOS
Figure 3.4 Synthetic molecules and xenobiotics acting as PPAR-α ligands
3.2.1.2 Endogenous ligands (biological molecules) of PPARα
The Gustafsson laboratory reported for the first time that endogenous fatty acids
can activate PPARα.12Interestingly, PPARα is the only subtype in the family which binds
to a variety of fatty acids with a high micromolar range of affinity.13 In human serum, the
concentration of free fatty acids is between 0.02-20 μM.14 Therefore, it is not well-
understood whether free fatty acid can be endogenous ligands for PPARα receptors. This
idea encouraged scientists to search for other molecules with interesting phenotypic
effects related to PPARs. In fact, they have discovered a number of oxidized metabolites,
such as eicosanoids, function as endogenous ligands in a tissue-specific manner. For
example, 8(S)-Hydroxy-(5Z,9E,11Z,14Z)-eicosatetraenoic acid(8-HETE) specifically activates PPARα.15 In addition, in a separate study on oxidized lipids, 9-hydroxy-
10E,12Z-octadecadienoic acid(9-HODE) and 13-hydroxy-9Z,11E-octadecadienoic 97
acid(13-HODE) derived from oxidized low-density lipoproteins (oxLDL) have shown
activation of PPARα.16 Another group interested in fatty acid catabolism working on hepatocytes has shown that when the fatty acid synthase (FAS) enzyme actively presents, a phospholipid, 1-palmitoyl-2-oleoyl-sn-glycerol-3-phosphocholine (16:0/18:1-GPC) acts as an endogenous ligand for PPARα.17 Recently, it has been demonstrated that in kupffer
cells, stimulation of the enzyme 5-lipoxygenase leads to activation of PPARα through production of intracellular 5S,12R-dihydroxy-6Z,8E,10E,14Z-eicosatetraenoic
18 acid(Leukotriene B4). (Figure 3.5)
OH COOH COOH
HO
8(S)-HETE 9-HODE
OH OH COOH COOH
OH
Leukotriene B4 13-HODE
O O P N O O O O O H
O
16:0/18:1-GPC
Figure 3.5 Endogenous (biological molecules) acting as PPAR-α ligands
3.2.2 Functional role of PPARγ
PPARγ has been highly studied among different species so far. It mostly plays a
key role in the differentiation of adipocyte tissues. It also modulates the expression of
genes involved in energy storage and utilization. The agonist of this receptor has been 98
demonstrated to improve the sensitivity of target tissues to insulin and to reduce plasma
glucose, lipid and insulin levels in type 2 diabetes mellitus.
3.2.2.1 Exogenous ligands of PPARγ
The major synthetic molecules for PPARγ belong to a family known as thiazolidinediones (TZDs) or “glitazones” used as antidiabetic agents. Their mechanism of action was not revealed until the mid-1990s, when their link to the differentiation of adipocyte tissues through PPARγ was proposed.19 Later, Spiegelman was able to show
that upon the binding of the ligand to PPARγ, the complex binds to the promoter region
of target genes.20 Two of the members of this family (rosiglitazone and troglitazone) are
shown in Figure 3.6. Since their discovery, extensive work has been performed to
expand the scaffolds targeting one or a combination of PPARs. Because of this, a number
of other drugs acting from nonrelated targets have shown activation of PPARγ at the
micromolar range, including LTD4 receptor antagonist LY171883 (tomelukast).
Furthermore, several nonsteroidal anti-inflammatory drugs (NSAIDs) such as
indometacin have shown PPARγ activation.21 (Figure3.6)
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S S O O N N NH O NH O O O O HO Rosiglitazone Troglitazone
O O OH O OH N N N O N N H O Cl
Tomelukast Indometacin
Figure 3.6 Synthetic molecules acting as PPARγ ligands
3.2.2.2 Endogenous ligands of PPARγ
Unlike the other subtypes, PPARγ has a strong preference for polyunsaturated acids. 13 PPARγ also binds to their oxidized metabolites from enzymatic or non- enzymatic reactions. For example, the lipoxygenase products of linoleic acid and arachidonic acid are 13(S)-HODE and 15(S)-HETE , respectively.22 In addition, the
12,14 prostaglandins 15-deoxy-Δ -prostaglandin J2 and 15-ketoprostaglandinE2 have shown activity at PPARγ.23,24 Furthermore, nitroalkene fatty acids 10-Nitrolinoleate and 9-
25,26 Nitrooleate have demonstrated PPARγ activity. (Figure 3.7)
100
COOH COOH
OH OH 13(S)-HODE 15(S)-HETE
O COOH COOH
O HO O ∆12,14 15-deoxy- J2 -Prostaglandin 15-keto Prostaglandin E2
NO2 COOH COOH
NO2 10-Nitrolinoleate 9-Nitrooleate
Figure 3.7 Endogenous molecules acting as PPARγ ligands
Phospholipids also exhibit affinity towards PPARγ. (Figure.3.8). Species of lysophosphatidic acid (LPA) such as 1-arachidonoyl-2-hydroxy-sn-glycero-3- phosphate(20:4 Lyso PA) have been proposed to act via PPARγ.27 However, LPAs
mediate their function mainly through G-protein coupled receptors on the cell surface. In addition, azelaoyl PAF derived from oxidized low-density lipoprotein (oxLDL) particles promote the differentiation of monocytes through the nuclear receptor PPARγ.28
(Figure.3.8)
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O O P O O OH O HO H 20:4 Lyso PA
O P N O O O O O H
O Azelaoyl PAF
Figure 3.8 Phospholipids which act as endogenous molecules acting as PPARγ ligands
3.2.3. Functional role of PPARβ/δ
PPARβ/δ has been identified as a regulatory factor in the development of several chronic diseases, such as obesity, atherosclerosis and cancer. Similar to PPARγ, activating PPARβ/δ can increase insulin sensitivity and improve glucose uptake in diabetic models. It is known that PPARβ/δ activates the expression of fatty acid- catabolizing enzymes in skeletal muscles. Research from the Evans laboratory has shown overexpression of this protein in mice muscles enhances physical strength and endurance.
In fact, these mice which are called “marathon mice” are able to run twice as far as normal mice. Although research on the development of drugs that can empower muscles is still under investigation, athletes have been reported abusing PPARβ/δ agonists to improve their performance in competitions. Consequently, the World Anti-Doping
Agency (WADA) has issued an alert to consumers about the health hazard of the available agonists on the black market.
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3.2.3.1 Exogenous ligands of PPARβ/δ
The initial efforts towards finding novel agonists for PPARβ/δ were mainly
focused on modification of natural fatty acids. For example L-631033 developed by
Merck had low micromolar binding affinity toward PPARβ/δ. Eventually, research
groups have found high-affinity ligands such as L-165041, GW501516 and GW 0742,
which have shown PPARβ/δ activation. (Figure.3.9)
COOH O COOH O
OH HO O O
O L-631033 L-165041
N N
F3C S F C S 3 S S F O COOH O COOH GW501516 GW0742
Figure 3.9 Synthetic molecules acting as PPARβ/δ ligands
3.2.3.2 Endogenous ligands of PPARβ/δ
It is known that PPARβ/δ exhibits a relatively high affinity for many saturated
13,29 and unsaturated fatty acids and eicosanoids. Eicosanoids PGA1, PGA2, PGD2, and
prostacyclin (PGI2) have been found to act as high affinity endogenous molecules towards the PPARβ/δ receptor.15,30,31 (Figure 3.10)
103
O O COOH COOH
OH OH PGA 1 PGA 2 COOH
OH O COOH
O HO OH OH PGD PGI 2 2
Figure 3.10 Endogenous molecules acting as PPARβ/δ ligands
3.3 EKODEs as potential endogenous metabolites
Although different research groups discovered a variety of fatty acids or related metabolites with high affinity towards PPARs, most of these findings were limited to a specific tissue. Therefore, there has been an interest towards finding endogenous metabolites which activate PPAR nuclear receptors in a global way.
So far, among different lipid metabolites that discussed in literature, the nitro form of fatty acid such as 10-Nitrolinoleate and 9-Nitrooleate can activate PPARγ subtype in human cells. In other words, the nitroalkene fatty acids which are products of nitrosative stress demonstrated a sub-micromolar range affinity to PPARs section 3.2.2.2.
These findings were quite important since they present how non-enzymatic pathways can regulate cell physiology in a concentration-dependent manner. Accordingly, our laboratory was sought the physiological role of lipid peroxidation products in cell
104
regulatory pathway feedback. In fact, the experiments designed here aimed to measure
the affinity of these molecules to the three PPAR nuclear receptors.
Among different molecules within the PUFA family that are known to be
modified during lipid peroxidation as a downstream of oxidative stress, we are focusing on molecules derived from linoleic acid called EKODEs. These molecules have shown interesting roles in regulating hormones, such as corticosterone and aldosterone production levels. They also demonstrate antioxidant effects through the Nrf2-Keap1 signaling pathway.32 In other words, the result from the Nrf2-Keap1 study was quite
remarkable because the products of peroxidation exhibit an anti-inflammatory effect by
triggering the detoxifying pathway of Nrf2. This idea raises the question whether the metabolites from this family can stimulate signals from other major known pathways
related to inflammation. In this regard, PPARs act as one of the major gene expression
machineries in the cell that are known to exhibit anti-inflammatory effects.33 As discussed
earlier, PPARs play a major role in regulating metabolism and catabolism in the body in
different tissues.
Along with other global techniques that are used to identify natural ligand for
PPAR, reporter assays still remain quite robust and reliable.34 In this study, a
heterologous expression setting of PPAR nuclear receptor in COS-7 cells was used. In
order to quantify the affinity of binding, a luciferase reporter system was employed. A
detail of fundamentals of the reporter assay will be presented in the following section.
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3.4 Target reporter assay
3.4.1 Bioluminescence
Bioluminescence has become more prevalent among researchers due to its
sensitivity and accuracy. In fact, luciferase-luciferin reaction provides a convenient reporter system to measure activities such as gene expression, post-translational modification (PTM), protein–protein interaction in a cell-based assay set up. Although fluorescence imaging techniques with the aid of green fluorescent proteins (GFPs) and synthetic dyes is still among the most common methods to study biological systems, the new discoveries in bioluminescence as an ultrasensitive approach is opening new doors to the field of cell physiology.
3.4.2 Cell-based reporter assay
Cell-based assays utilizing reporter enzymes are widely used widely to investigate promoters, interaction between promoters and transcription factors, signal transduction, and other cellular activities. In fact, they provide great advantages such as being rapid and relatively inexpensive making them ideal for drug screening to study drug targets and signaling pathways. Among the reporter genes, luciferase, an enzyme which catalyzes the bioluminescence reaction has been used more frequently. In fact, comparing to the other typical enzymes such as chloramphenicol acetyltransferase, β-glucuronidase, luciferase is highly sensitive and it gives a linear response over at least eight orders of magnitude. In addition, another advantage of this method is the ease of measuring light intensity with photomultiplier of a charge-coupled device (CCD) camera. Accordingly, luciferase is a suitable reporter system for measuring gene expression quantitavily.35
106
3.4.3 Single luciferase reporter assay
All types of bioluminescence luciferase systems follow the same principle of
chemical reaction: at the end the electronically exited state product will release light as a bioluminescence signal. The high emission intensity depends on the extraordinary quantum yield of the reaction (ΦBL). Indeed, the quantum yield in the case of the firefly
luciferase reaction is >0.40 which means that four photons can be obtained from ten
reacting molecules.36 In addition, the intensity of light is dependent on the amount of
reagents (ATP, Mg2+, and oxygen) and concentration of the enzyme. In order to measure
transactivation of PPAR nuclear receptors with their ligand, the response element (PPRE)
attached to a luciferase gene in the reporter vector along with another vector containing
the PPAR receptor gene are transfected into target cells.
Luciferase Living cell Ligand PPRE reporter gene
PPAR transfection
βGal gene
PPAR
transfection
PPAR gene
mRNA transfection
Luciferase protein
βGal protein
PPAR
Luciferin
cell extract
Figure 3.11 Illustration of the basic principle of the single luciferase reporter assay for PPAR receptor
107
Because luciferase expression is controlled by the fused genetic elements, reporter
expression is directly correlated with the activity of the PPRE. After transfection of the
plasmid into target cells, the PPAR vector provides the protein, which can interact with
the ligand and eventually move toward the PPRE. In other words, transactivation of
PPARs with the ligand leads to expression of luciferase and emitting the bioluminescence signal. The amount of signal is a good estimation of the activity of luciferase protein, which indicates the binding affinity of ligand to the PPAR nuclear receptor in the cells.
3.4.4 Luciferase enzyme mechanism
Luciferase acts as an enzyme and in the presence of cofactors (ATP, Mg2+ and
oxygen) it conducts an oxidative reaction on the substrate luciferin. At the beginning it
proceeds as an acyl-adenylate intermediate from a carboxylic acid substrate and ATP.
Next, luciferase as a monooxygenase adds molecular oxygen to luciferyl acyl-adenylate
complex creating a highly strained dioxentanone. At this stage the enzyme promotes
cleavage of CO2, forming oxyluciferin through [2+2] retero-cycloaddition, which is
forbidden by orbital symmetry law.37 Then the electronic excited state emits light while
returning to the ground state.38,39 As a result, this method can give a quantitative
measurement of ligand-PPAR interaction.40
108
O N N COOH MgATP PPi N N AMP O2 O S S luciferase O S S D-Luciferin D-Luciferyl Adenylate
O O O O AMP N N N N O O O S S O S S AMP CO2 dioxentanone * O N N N N O S O S S O S
excited oxyluciferin Light
Figure 3.12 Reaction catalyzed by firefly luciferase emits lights through bioluminescence process
3.4.5 Colorimetric β-galactosidase assay
To eliminate intrinsic experimental error, such as differences in the number and
viability of cells used, as well as the efficacy of the cell transfection, and lysis, etc. The
beta-galactosidase (βgal) gene is transfected to the cells in a separate vector at the same time. This enzyme is resistant to protease degradation and is stable in physiological condition. The enzyme which is the product of LacZ gene recognizes terminal nonreducing β-D-galactose residue in β -D-galactopyranosides. For example, hydrolysis of lactose provides β -D-galactose and glucose. Interestingly the enzyme is capable of hydrolysis of a broad range of substrates as long as they possess the β-D-galactose 109
residue. As a result, scientists took advantage of this property and designed substrates
that eventually broke into colored products. One of the widely used substrates is the ortho-Nitrophenyl-β-galactoside (ONPG) molecule which can be hydrolyzed to β -D-
galactose and ortho-nitrophenol(ONP). The latter absorbs light at 420 nm (seen as intense
yellow color) while the precursor molecule ONPG does not. Thus, by using ONPG as a
substrate, the activity of the enzyme can be quantified by measuring the amount of
absorption of ONP at 420 nm using a spectrophotometer or a microplate reader.
OH OH OH NO OH H O 2 NO2 H H O O β HO HO -gal H OH H OH HO + H H H OH H H
ONPG β-D-galactose ONP
chromophore
Figure3.13 Colorimetric assay of β-galactosidase using ONPG as the substrate
110
3.5 Experimental
3.5.1 Materials
Each compound was prepared in DMSO at 1000x prior the experiment.
Specifically, compounds and concentrations were as follows: 4-Chloro-6-(2,3-xylidino)-
2-pyrimidinylthioacetic acid (Wy14643; Sigma) prepared at 5mM concentration, 4-[2-(3-
Fluoro-4-trifluoromethyl-phenyl)-4-methyl-thiazol-5-ylmethylsulfanyl]- 2-methyl- phenoxy}-acetic acid(GW0742; Sigma) prepared at 5mM concentration, 5-[4-[(6-
Hydroxy-2,5,7,8-tetramethylchroman-2-yl)methoxy]benzyl]-2,4-thiazolidinedione,
(troglitazone )prepared at 5mM linoleic acid 4-hydroxynonenal(4-HNE), 4- oxononenal(4-ONE), 10-nitrolinoleate, 10(E),12(Z)-conjugated linoleic (Cayman
Chemical). EKODE molecules were synthesized, as described in Chapter 2, and purified by HPLC. EKODE amount were gravimetrically quantified using an AD-6 microbalance
(Perkin Elmer Corporation, Norwich, CT). All the lipids and oxidized metabolites stock solutions were prepared at 20.0mM. Cell Culture Dishes were 100mmx20mm, 12-well plate (Corning).
3.5.2 Cell transient transfection assay
COS-7 cells from American Type Culture Collection (ATCC) were grown to 60% confluence in Dubelco's Modified Eagle Medium (DMEM, Corning, Manassas, VA) supplemented with 10% FBS and 1% Antibiotic-Antimycotic (Life Technologies). Cells
were incubated at 37°C and 5% CO2 in a Forma™ Series II 3110 Water-Jacketed CO incubator (Thermo Scientific). Then COS-7 cells were transiently transfected with a plasmid containing the luciferase gene under the control of three tandem PPAR response elements (PPRE) (PPRE X3 -TK-luciferase) and mousePPARα, mousePPARγ, or
111 mousePPARβ/δ expression plasmids(Addgene, Cambridge, MA), respectively. pSV-β-
Galactosidase plasmid (Promega, Madison,WI) was co-transfected to monitor transfection efficiency. Plasmids were transiently transfected using PolyFect (Qiagen
301107). Cell lysates were prepared and luciferase and β-galactosidase activities were assayed using kits as described in methods section 3.5.2.5. Readings were taken with a
Flex Station 3 plate reader (Molecular devices) (Pharmacology department, Case Western
Reserve University).
3.5.3 Methods
3.5.3.1 Seeding of COS-7 cells
10 5 cells were seeded to each well in a 12-well plate in DMEM (supplemented with 10% FBS, 1% Antibiotic-Antimycotic streptomycin) as described in 3.6.1.
3.5.3.2 Transfection of COS-7 cells
After 24 hours of seeding and letting cells grow to 60% confluence, the growth media was replaced with a fresh media and transfection occurred. Cells were transfected with 200 ng PPRE X3 -TK-luciferase, 20 ng PPAR, PPAR, PPARγ, 100 ng internal reporter plasmid pSV-β-Galactosidaseand. A mix of these plasmid constructs and 4 μL
Polyfect reagent reached to the final volume of 40 μL with charcoal-treated DMEM.
Transfection mixture was incubated at room temperature for 10 minutes. Adding 40 μL of the media (DMEM, supplemented with 10% FBS, 1% Antibiotic-Antimycotic streptomycin) to the mixture brought the volume to 80 μL, which was dispensed into each well. Transfected cells were incubated for overnight prior to the treatment.
112
3.5.2.3 Treatment of transfected cells
The growth media was replaced with a charcoal-treated media for 4 hours.
Subsequently 1 μL of the ligands from 1000x stock solution in DMSO was added to each well. For the control experiment, activity of vehicle (DMSO) was tested by adding 1 μL
to each well. Ligand-treated cells were incubated for 24 hours prior to the readout.
3.5.2.4 Cell lysis
The media were removed from each well and cells washed once with 1 x PBS.
Then a 150 μL lysis buffer was added to each well. After scraping the cell remnants,
plates were kept at 4oC.
3.5.2.5 Luciferase assay, data normalization
In order to measure the luciferase activity in ligand- treated cells, the manufacturer protocol was followed. From each well 20 μL of cell lysate was placed in a
96-well opaque plate (white bottom) and treated with 100 μL of luciferase mixture.
(Caution: the addition needs to be quick to eliminate the error in signal collection). In order to normalize the data, internal reporter plasmid pSV-β-Galactosidase was used.
Following the instructions of the pSV-β-Galactosidase manufacturer, 30 μL of cell lysate from each well was placed in a 96-well plate, and a mixture containing ONPG added.
After one hour of incubation at 37 oC, the signal measured at 420 nm. Data were
collected as the relative light units (RLU) obtained from the luciferase assay divided by
the corresponding absorbance value obtained at 420 nm in the β-galactosidase assays
(Luc/β-galactosidase). Each compound was repeated in triplicate in a single experiment.
Therefore, values from the replicate were then averaged and expressed as mean normalized firefly luciferase activity.
113
3.6 Result and discussion
As we discussed earlier, transactivation of PPAR receptors with ligands, activates
luciferase expression and the signal obtained can be used as a quantitative measurement
for affinity of the ligand to PPAR. The result shown in Figure 3.14 is the luciferase read
out normalized by βgal which was used as internal reporter assay. In the case of
mPPARα, synthetic ligand Wy14635 was used at 5 μM. Among different lipids and
related metabolites of EKODEs, trans-EKODE-(E)-IIa and trans-EKODE-(E)-IIb showed a better activity, however this activation is not comparable to the synthetic ligands.
Figure3.14 Relative light units (RLU) determines activity of PUFAs and lipid metabolites on mouse PPARα by luciferase assay. 1X(5μM), 2X(10μM), 4X(20μM). Wy14635 is the synthetic ligand
114
Similarly the data for mPPARγ presented in Figure 3.15. Among different lipids and related metabolites of EKODE, trans-EKODE-(E)-IIa and trans-EKODE-(E)-IIb show a slightly better activity.
Figure3.15 Relative light units (RLU) determines activity of PUFAs and lipid metabolites on mouse PPARγ by luciferase assay. 1X(5μM), 2X(10μM), 4X(20μM). Troglitazone is the synthetic ligand
Interestingly, trans-EKODE-(E)-IIb showed a tremendous activation towards mPPARβ/δ which can be interpreted as specific interaction of this metabolite with mPPARβ/δ receptor. In this case activation by 10 μM of this metabolite is comparable with GW0742
at 5μM Figure 3.15.
115
Figure3.16 Relative light units (RLU) determines activity of PUFAs and lipid metabolites on mouse PPARβ/δ by luciferase assay. 1X(5μM), 2X(10μM), 4X(20μM). GW0742 is the synthetic ligand
In order to understand this exceptional activation, another experiment with lower
concentration of trans-EKODE-(E)-IIb was performed. Interestingly, this metabolite shows activity in a dose-response manner between 2.5-20 μM of concentration Figures
3.16 & 3.17.
116
Figure3.17 Relative light units (RLU) determines trans-EKODE-(E)-IIb activation of mouse
PPARβ/δ in compare to GW0742
3.7 Conclusion
Since the PPAR nuclear receptors has been intervened in different metabolic
networks and related diseases. Finding a high affinity endogenous ligand can shed light
on the PPAR role in the maintenance of cell physiology and therapeutics for variety of
diseases.41 Interestingly, nitrosative products of fatty acids have been known as
endogenous ligand for PPARγ, which emphasizes on the fact that downstream signaling
of reactive nitrogen species (RNS) can be important in the maintaining of cell
physiology. In the case of nitration products, they exhibit phenotypic effect as it has been
observed by PPARγ activation through synthetic ligands. Indeed, this observation along
with binding studies to the receptor suggested that the nitrated fatty acids function
through PPARγ.25
Similarly our result indicates that trans-EKODE-(E)-IIb activates PPARβ/δ in a dose dependent manner provides the evidence that this molecule can be nominated as 117
possible endogenous activator for PPARβ/δ. However, more detail studies need to be
performed in order to reveal their mode of interaction. The outcome of this finding will
definitely have a great impact on understanding of diseases which are associated with
PPARβ/δ such as chronic metabolic diseases and muscle dystrophy. In addition, at this
point designing specific molecules with the core structure of trans-EKODE-(E)-II can be found beneficial.
118
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Chapter 4 Thesis summary and future directions
4.1 Thesis summary
In Chapter 1, the discovery of oxidized lipids as endogenous metabolites
has been discussed. First, prostaglandins (PGs) and leukotrienes (LT) with a short
overview of their history were introduced. The role of stereocontrolled synthesis
in characterization and understanding of the biosynthetic pathways associated
with these metabolites was introduced. Next, the discovery of lipid peroxidation
products (LPO) derived from non-enzymatic reactions was discussed. The
synthesis of LPOs along with their physiological significance was then introduced. The interplay between oxidative stress and modification of lipids was deliberated. Next, the background of oxidative stress was introduced. Based on a
comprehensive literature review, the different types of ROS/RNS were discussed
and how their physicochemical properties can determine their reaction kinetics
and thermodynamics. The discussion continued with the chemical basis of how
lipid peroxidation products (LPO) can activate diverse biotargets.
In Chapter 2, one of the main products of linoleic acid peroxidation, or
EKODEs, was introduced. These molecules have been found in pathological
states with an oxidative stress background. Following, the postulate mechanism
was shown regarding how ROS/RNS initiates radical formation, and bonds
rearrange themselves to form a variety of scaffolds. The fact that these molecules
possess an α, β-unsaturated ketone structure makes them more interesting to
investigate to define their role in cell physiology. The detailed synthesis for
123
different combinations of this family of metabolites was also discussed. Our
challenges during the synthesis of EKODE-(E)-II encouraged us to implement a novel strategy through a divergent approach to synthesize these metabolites and related molecules. In other words, we have shown how this proposed synthetic strategy can help us to build different libraries of EKODE-like molecules. The fact that we were able to design a core reactive “bifunctional conjunctive ylide”
22 helped us to obtain these metabolites in high yields in a rapid timeframe.
O
Ph3P S(CH3)2 22
Bifunctional conjunctive ylide
RCHO RCHO
k2 k1
O O Ph3P R S(CH3)2 R O
Scheme.4.1 At low temperature (0oC), sulfonium ylide reacts faster than phosphonium
(k1>>k2)
As shown in Scheme 4.1 the bifunctional molecule takes advantage of the
difference in reactivity between two ylides attached to the main scaffold of
EKODE-(E)-II. The importance of this methodology is that the ylides exhibit
different nucleophilicity while reacting with aldehydes providing an ideal
opportunity for substrate discrimination based on inherent reactivity. In this
example, the sulfonium ylide reacts faster than phosphonium, providing an
advantage when performing sequential reactions with the aldehydes of interest.
124
O O 1eq R1CHO 1eq R2CHO Ph P 3 S(CH3)2 R R Epoxide formation Olefin formation 2 O 1 22
Bifunctional conjunctive ylide EKODE-(E)-II
Scheme.4.2 Sulfonium ylide reaction is faster than phosphonium one
The use of the bifunctional conjunctive ylide resulted in production of EKODE-
(E)-II and their alkyne-terminal bioorthogonal forms and 2H-labeled derivatives that can be used for further mechanistic studies. The strategy outlined here establishes a general approach that can be applied to a wide range of PUFA metabolites, to develop libraries of reactive lipid metabolites to study their possible biological functions.
In Chapter 3, the biological evaluation of PPAR nuclear receptors was performed. The previous kinetic studies on these metabolites, along with other end products of LPO, suggested that they are acting through formation of Michael adducts in a comparable rate.1 The result provided a stepping-stone toward our
ambition of revealing biological targets for these metabolites.
A luciferase reporter assay was utilized to measure the transactivation of
these lipid metabolites toward PPAR nuclear receptors. In a heterologous
expression setting of PPAR nuclear receptor in COS-7 cells, transactivation of the
PPAR nuclear receptor with the respective ligands was measured. Interesting
results were observed showing trans-EKODE-(E)-IIb activating PPARβ/δ nuclear receptors. These receptors are specifically involved in the development of different chronic diseases, such as obesity, atherosclerosis and cancer.The result from this assay indicated that this specific metabolite can regulate cell signaling
125
through PPARβ/δ. However, the results here raise a fundamental question
regarding whether these molecules can regulate chronic diseases, which obviously
needs further investigation. In addition, the detailed of the mode of interaction needs to be revealed via more thorough structural studies, such as X-ray crystallography.
4.2 Future directions
To facilitate a deeper understanding of the function of these metabolites, our future investigation is going to follow on two parallel directions: first, our laboratory is currently designing a set of global experiments such as RNA-seq or
microarray experiments to characterize functional targets in the cell. The outcome of this experiment will be critical in order to provide us with preliminary data.
Consequently, the specific targets will be confirmed through RT-PCR experiments, and quantification of RNA expression levels will be used to evaluate data obtained from global studies. The information will help us build models for proposing networks regulated by EKODEs. In addition, understanding how these specific metabolites are interacting with proteins on metabolic or signaling networks will provide us deeper insight toward revealing their mechanisms of action. By understanding how these metabolites are interacting with cellular lipase enzymes and intracellular carrier proteins such as fatty acid binding proteins (FABPs), research can determine how these interactions can affect signaling and the metabolism of other fatty acids. In this regard, the bioorthogonal surrogate and deuterated-labeled molecules, which were developed through our
126
novel synthetic approach, will likely be very beneficial. Structural studies such as
X-ray crystallography can be employed to investigate the interaction modes at the
level of molecular detail.
Our second focus will be on employing the synthetic strategy represented
in this thesis for developing a variety of related molecules from other PUFAs and
investigating their biological implications. In addition, these oxidized fatty acids
can be conjugated to the structure of phospholipids such as phosphatidylcholine
and lysophosphatidic acids, which have shown interesting results in activating G-
protein coupled receptors (GPCR) and nuclear receptors with other oxidized
lipids.2-4 Synthesis of phospholipids can be achieved with different commercially
available phosphorous containing heads through synthetic or chemoenzymatic
esterification as shown in Scheme 4.3.5-7
O OR OR esterification Oxidized fatty acids O Oxidized fatty acids COOH + HO Phsophorous Phsophorous head head
R: acylgroup or H Scheme 4.3 General strategy for phospholipid synthesis
Overall, the versatile synthetic approach which has been presented in this thesis
provides the synthesis of the EKODE family and similar analogues to facilitate
the biological evaluation for the future.
127
4.3 References
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APPENDIX NMR spectra of synthesized molecules
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135
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