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University of Southern California Dissertations and Theses

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Studies on lipid mediators, and on potential modulators of GRP78

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Studies on lipid mediators, and on potential modulators of GRP78

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Content i

STUDIES ON LIPID MEDIATORS, AND ON
POTENTIAL MODULATORS OF GRP78

by

Anne-Marie Finaldi

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(CHEMISTRY)

December 2013

Copyright 2013 Anne-Marie Finaldi

ii
Dedication

This work is dedicated to my father, Umberto Finaldi

iii
Acknowledgments

Above all, I would like to thank Professor Nicos A. Petasis for all of his guidance,
support, and encouragement throughout my graduate career. Without his help and wisdom none
of this work would have been possible. It is extremely rare to meet someone as patient,
understanding, knowledgeable, and creative as Professor Petasis and I am honored to have been
under his advisement.
I would also like to express my sincerest gratitude to Professor G. K. Surya Prakash for
his kindness, advice, and help along my journey. His brilliance and humbleness have been an
inspiration to me, and I feel privileged to know him.
I wish to thank my committee members Professors: Barry C. Thompson, Mark E.
Thompson, and Stan Louie for their encouragement and helpful discussions.
Special thanks go to Professor Charles E. McKenna and Dr. Boris Kashemirov for the use
of their LC-MS/MS/MS instrument and technical advice. I would also like to thank Professor
Amy S. Lee for her collaboration and support.
I would also like to thank Carole Phillips, David Hunter, and Michele Dea for their
consistently proficient work and for always helping me. My sincerest thanks also go to Dr. James
Ellern and Dr. Jennifer Moore, for their guidance, support and kindness during my teaching
assistantship.
I am grateful to my past and present group members, specifically: Dr. Rong Yang, Dr.
Jeffery C. Raber, Dr. Kalyan Nagulapalli, Dr. Malgorzata Myslinska, Dr. Alexey Butkevich,
Dr. Jamie Jarusiewicz, Min Zhu, Charles Arden, Marcos Sainz, Stephen Glynn, and Michael
Carlson for their friendship, intellectual discussions, and for teaching me patience.
iv
I cannot express how thankful I am for my friends who have listened to me, nurtured me,
and strengthened me during my graduate studies at USC. Notably, Dr. Fang Wang, Dr. Ivan
Krylov, Valentina Krylova, Anton Shakmin, Dr. Parag Jog, Hema Krishnan, Andrey Rudenko,
Richard Giles, Dr. Mikhail Zibinsky, Dr. Na Zhang Cao, and Dr. Tangyuan Cao were an integral
part of my success and I cannot thank them enough for the friendship and kindness they have
shown me.
I am immensely grateful to Sushma Misra, Ram Misra, and Shivani Misra, for their
hospitality, compassion, and generosity they have shown me. It is truly rare to meet such kind-
hearted, warm, and caring people and I am lucky to know them.
I would like to give a special thanks to my family, specifically Leonardo Finaldi, Diana
Finaldi, Diana Dyke, Ashley Dyke, and David Dyke, for their endless support and love.
Leonardo has been such an amazing role model to me growing up and his encouragement and
humor have gotten me through some of my major challenges in graduate school. From the
bottom of my heart I want to thank my late father, Umberto Finaldi, for being the best father any
person could ask for. My father has been very inspirational to me, showing me that even when
things are painful and difficult, we should never lose hope.
Finally, I would like to thank Nitin Misra for all of his infinite love and support. His
patience, intelligence, sensitivity, and loyalty never cease to amaze me. No one else has ever
understood me or cared for me the way that Nitin has, and to him I am extremely grateful.

v
Table of Contents
Dedication ii
Acknowledgements iii
List of Figures vii
List of Schemes viii
Abstract x
Chapter 1. Polyunsaturated Fatty Acids, Lipid Mediators, and Inflammation 1
1.1 Introduction 1
1.2 The Lipoxygenation Pathways in Inflammation 2
1.3 DHA-Derived Lipid Mediators 6
1.4 AA-Derived Lipid Mediators 8
1.5 Conclusion 10
1.6 Chapter 1 References 12

Chapter 2. A Docosahexaenoic Acid Analog as an Imaging Probe for Lipid Mediator
Pathways 13
2.1 Introduction 13
2.2 Results and Discussion 18
2.2.1 Design of an ω-Alkynyl-DHA Derivative 18
2.2.2 First Total Synthesis of an ω-Alkynyl-DHA Derivative as an Imaging
Probe 19
2.2.3 Lipoxygenation of an ω-Alkynyl-DHA Derivative with
15-Lipoxygenase 23
2.3 Conclusion 27
2.4 Experimental 27
2.5 Chapter 2 References 35

Chapter 3. Design and Synthesis of Lipoxin Analog Building Blocks 37
3.1 Introduction 37
3.2 Results and Discussion 39
3.2.1 Design of Lipoxin Analog Building Blocks 39
3.2.2 Synthesis of Fluorophenoxy Building Blocks 42
3.2.3 Synthesis of 3-Oxa Lipoxin A
4
Analog Building Blocks 44
3.2.4 Synthesis of Benzo-Lipoxin Building Blocks 49
3.3 Conclusion 51
vi
3.4 Experimental 51
3.5 Chapter 3 References 70

Chapter 4. Synthesis of Potential Modulators of GRP78 72
4.1 Introduction 72
4.2 Results and Discussion 75
4.2.1 Design of a Small Library of Compounds for testing GRP78 Activity 75
4.2.2 Retrosynthesis of Potential Modulators of GRP78 78
4.2.3 Synthesis of Potential GRP78 Modulator Building Blocks 80
4.2.4 Assembly of Building Blocks into Final Products 85
4.3 Conclusion 90
4.4 Experimental 90
4.5 Chapter 4 References 107

Conclusions 109

Bibliography 110

Appendix: Selected Spectra 115

vii
List of Figures
Figure 1.1 The metabolism of fatty acids into pro- and anti-inflammatory
lipid mediators upon infection or injury 2
Figure 1.2 The anti-inflammatory and pro-resolving action of lipid mediators
generated by 15-lipoxygenase 5
Figure 1.3 Arachidonic acid and the lipid mediators it is metabolized into 9
Figure 2.1 Bertozzi’s copper-free click chemistry in living animals 17
Figure 2.2 A novel DHA analog as an imaging probe for lipid mediator pathways 23
Figure 3.1 Structures of potent anti-inflammatory agents, lipoxins (LXA
4
and
LXB
4
) and aspirin-triggered lipoxins (15-epi-LXA
4
and 15-epi-LXB
4
) 37
Figure 3.2 Select lipoxin analogs that have exhibited high anti-inflammatory
potency and enhanced metabolic stability 38
Figure 3.3 Retrosynthesis of para-fluorophenoxy building blocks 3.1 and 3.2
from a chiral starting material 40
Figure 3.4 Retrosynthesis of 3-oxa lipoxin A
4
building blocks 3.3 and 3.4,
derived from the chiral sugar L-Rhamnoser 41
Figure 4.1 Stress-induced GRP78 activity within a tumor cell 74
Figure 4.2 The design of novel hexahydropyrazinotriazinedione β-turn
peptidomimetics for the modulation of GRP78 77
Figure 4.3 Structures of novel final compounds for potential GRP78 modulation
synthesized herein 78
Figure 4.4 Essential carboxylic acid building blocks and amine building
blocks for the synthesis of the the title compounds 80

viii
List of Schemes

Scheme 1.1 Biosynthesis of neuroprotectin D1 (NPD1) from DHA 6
Scheme 1.2 Aspirin-triggered resolvin D-series compounds biosynthesized from DHA 8
Scheme 2.1 DHA and a few of the lipid mediators it is metabolized into
(resolvins D1 and D2, neuroprotectin D1, and maresin R1) 13
Scheme 2.2 A general example of a “click” reaction 15
Scheme 2.3 Click chemistry that results in a fluorescent product 16
Scheme 2.4 A retrosynthetic analysis of the alkynyl-DHA derivative target 19
Scheme 2.5 The first half of the total synthesis of ω-alkynyl-DHA (2.1),
a potential imaging probe for lipid mediator pathways 20
Scheme 2.6 The second half of the total synthesis of the title compound (2.1) 22
Scheme 2.7 Lipoxygenation reaction with 15-lipoxygenase (15-LO) and
standard DHA or ω-alkynyl-DHA 26
Scheme 3.1 The first part of the synthesis of para-fluorophenoxy building
blocks 3.1 and 3.2 42
Scheme 3.2 The second half of the synthesis of vinyl iodide (3.1) and
alkyne (3.2) para-fluorophenoxy building blocks 43
Scheme 3.3 The synthesis of 3-oxa vinyl iodide building block 3.3 45
Scheme 3.4 The synthesis of 3-oxa vinyl iodide 3.4, similar to the synthesis
of 3.3, but with different protecting groups 46
Scheme 3.5 A Wittig reaction to couple the aldehyde (3.14) with the phosphonium
salt (3.19) to produce 3.20, and the attempted isomerization of the
cis/trans mixture (3.20) to 3.21 48
Scheme 3.6 Suzuki coupling and Miyaura borylation reactions to produce the
meta-benzo-lipoxin building blocks 49
Scheme 3.7 Suzuki coupling and Miyaura borylation reactions to produce the
para-benzo-lipoxin building blocks 50
Scheme 4.1 The retrosynthesis of hexahydropyrazinotriazinedione final
compounds from a peptide-like backbone structure 78
ix
Scheme 4.2 The retrosynthesis of 4.4, the general peptide-like backbone 79
Scheme 4.3 The synthesis of carboxylic acid 4.7 in four overall steps 81
Scheme 4.4 The synthesis of carboxylic acid 4.8 utilizing a reductive amination
in place of the SN
2
reaction used for the synthesis of 4.7 83
Scheme 4.5 The synthesis of amine building block 4.9 in three steps 84
Scheme 4.6 The synthesis of 4.10 using a peptide coupling reaction and a deprotection 84
Scheme 4.7 The synthesis of amine 4.11 in two steps 85
Scheme 4.8 Peptide coupling to produce 4.22, the precursor of final product 4.1 86
Scheme 4.9 The synthesis of 4.23 using HATU as a peptide coupling reagent 87
Scheme 4.10 The synthesis of backbone 4.24 to be later cyclized into 4.3 87
Scheme 4.11 The final step in the synthesis of final product 4.1 88
Scheme 4.12 The final step in the synthesis of final product 4.2 89
Scheme 4.13 The final step in the synthesis of final product 4.3 89

x
Abstract

The research presented within this dissertation is a compilation of three projects with the
underlying theme of designing and synthesizing relevant molecules for biological studies.
The first chapter serves as an introduction to lipid mediators and their role in
inflammation. A brief overview of the lipid mediator pathways and their significance are
discussed within chapter one.
Chapter two introduces click chemistry as a tool for probing lipid-related biological
pathways. The design and total synthesis of an alkynyl-docosahexaenoic acid analog for
potential use as an imagining probe using click chemistry is described. Preliminary enzymatic
assays were also performed to assess the stability and metabolism of the final product.
The third chapter presents the design and synthesis of building blocks for the total
synthesis of lipoxin analogs. The building block design was focused on incorporating moieties
that would potentially be metabolically stable and biologically active upon later completion of
the final structure.
Chapter four introduces the protein GRP78 and its significance to cancer and disease
development. The concept of reverse-turn peptidomimetics is briefly introduced and applied to
the design of our target molecules. Novel potential modulators of GRP78 are successfully
synthesized for biological testing.

1
Chapter 1. Polyunsaturated Fatty Acids, Lipid Mediators, and
Inflammation
1.1 Introduction
Inflammation is the body’s natural response to protect itself from injury, infection, and
other harmful invasions. However, when inflammation is persistant, such as in a chronic
inflammatory disease, the harmful effects on the body can be more than just painful.
Inflammation has been linked to several major diseases such as cancer, Alzheimer’s Disease,
arthritis, and heart disease. Furthermore, head injury results in inflammation and without proper
resolution of the inflammation, adverse effects such as brain damage can occur. Understanding
inflammatory signaling pathways and resolution is of great importance because it could give rise
to more effective treatments for diseases that are currently incurable. It has long been known that
polyunsaturated fatty acids (PUFAs) such as docosahexaenoic acid (DHA), eicosapentanoic acid
(EPA), and arachidonic acid (AA) play a critical role in inflammation. Although it is well
established that omega-3 fatty acids (such as DHA and EPA) are beneficial to resolving
inflammation, their mechanism of action has only recently begun to be elucidated.
1

The significance of polyunsaturated fatty acids in development and health was discovered
by Burr and Burr in 1929, when they found that a rat’s diet devoid of essential fatty acids
resulted in disease and premature death.
2
While this work was intriguing, there were still a
number of unanswered questions regarding which fatty acids were essential, and the mechanism
in which they were acting through. Later in 1975, Dyerberg, Bang and Hjorne discovered that
the Eskimos of Greenland who consumed a diet high in PUFAs from fish oil had a much lower
incidence of heart disease than people from industrialized countries.
3
These findings were highly
2
influential in the field of lipids, as it furthered the connection between the consumption of
PUFAs and inflammatory disease mitigation. The discovery of the prostaglandins and
leukotrienes, derived from the omega-6 PUFA arachidonic acid (AA), revealed the role of
certain PUFA metabolites to promote inflammation, and serve as proinflammatory lipid
mediators. Lipid mediators are the signaling molecules derived from lipids, which control the
inflammatory pathways. In recent years several classes of anti-inflammatory and pro-resolving
lipid mediators were identified (lipoxins, resolvins, protectins, maresins, etc.), and within each
class there are several mediators with well-established structure and activity profile. However,
new lipid mediators are being discovered and are of particular interest because of their high
importance to inflammation.

1.2 The Lipoxygenation Pathways in Inflammation

Figure 1.1- The metabolism of fatty acids into pro- and anti-inflammatory lipid mediators upon
infection or injury.
1, 4

3
Omega-3 fatty acids are polyunsaturated fatty acids that contain a double bond three
carbons away from the omega (terminal) end of the fatty acid. Omega-6 fatty acids contain a
double bond six carbons away from their terminal end. Omega-6 fatty acids trigger pro-
inflammatory responses, while omega-3 fatty acids have been found to have anti-flammatory
properties. Examples of omega-3 (EPA and DHA) and omega-6 (AA) fatty acids are shown in
Figure 1.1.
1, 4
The balance between pro- and anti-inflammatory responses is very critical to
healthy function.
Humans get many of their omega-3 and omega-6 fatty acids from digesting the fish,
meats, and other foods that we consume. These fatty acids become a part of the phospholipid
bilayer membrane of our cells, but when inflammation occurs phospholipase enzymes cleave
specific bonds that release the fatty acids so that they may initiate inflammatory pathway
signaling.
1
Oxygenating enzymes, such as cyclooxygenases (COX) and lipoxygenases (LO),
then metabolize the free fatty acids. The lipids can be directly synthesized into a lipid mediator
by interacting with an enzyme once, or they can be subjected to a cascade mechanism in which
the lipid is sequentially transformed by the same or other enzymes. Figure 1.1 shows a broad
overview of some of lipid metabolism pathways that result in critical lipid mediators of
inflammation.
Lipoxygenases are enzymes that use the iron in their active site to stereoselectively
oxygenate specific positions on PUFAs.
5, 6
15-Lipoxygenase (15-LO) is one type of
lipoxygenase that is extremely important to inflammation resolution, so named because it
oxygenates arachidonate at the 15-carbon position. Among the common lipoxygenases currently
known (5-, 12-, and 15-), 15-LO is recognized for producing the widest range of anti-
inflammatory lipid mediators. 15-LO is responsible for the production of lipoxin A
4
(LXA4),
4
resolvins D1 and D2 (RvD1 and RvD2), neuroprotectin D1 (NPD1), and maresin 1 (MaR1)
(Figure 1.1).
1
All of these lipid mediators are essential to brain and eye function, as well as
general health and well-being.

15-Lipoxygenase is classified into two different isoforms, 15-lipoxygenase-1 (15-LOX-1,
also called 12/15-lipoxygenase and 12/15-LO) and 15-lipoxygenase-2 (15-LOX-2).
7
15-LOX-1
is the more abundant and diverse isoform, expressed in several various tissue types all over the
body, and with the enzymatic activity of 12-lipoxygenase as well.
6
15-lipoxygenase-2 is
considered more specialized, as it is mainly expressed in prostate and skin cells.
6
Both isoforms
of 15-lipoxygenase have been linked to prostate cancer, with evidence pointing to 15-LOX-1 and
15-LOX-2 having counteracting effects, although more data is needed to thoroughly elucidate
their roles in prostate cancer regulation.
8,

9
A recent clinical trial showed that in breast cancer the
ratio of 15-LOX-1 to 15-LOX-2 expression was predictive of the medical outcome of the
patient.
6
Furthermore, both 15-LOX-1 and 15-LOX-2 were found to suppress tumor growth in
breast cancer patients.
6
Even though the precise role of 15-LOX-1 in cancer and other
inflammatory diseases is not clearly understood, it is certain that the activity of 15-LOX-1 must
be strictly regulated for proper cell function and development.
7
15-LOX has several actions, as well as synthesizing lipid mediators directly from PUFAs,
it also converts the intermediate leukotriene A
4
produced by 5-LOX into LXA
4
.
10
Unlike 5-
lipoxygenase, which synthesizes both pro- and anti-inflammatory lipid mediators, 15-
lipoxygenase only produces anti-inflammatory lipid mediators. Interestingly, pro-inflammatory
lipid mediators prostaglandins E
2
and D
2
activate 15-LOX expression, which in turn causes 15-
LOX to produce more LXA
4
and this leads to inflammation resolution.
11, 10

5

Figure 1.2- The anti-inflammatory and pro-resolving action of lipid mediators generated by 15-
lipoxygenase.
12

In inflammation, 15-lipoxygenase produces lipid mediators that actively enhance the
resolution of inflammation (see Figure 1.2). Certain lipoxins, resolvins, and protectins
synthesized by 15-LOX-1 have a two-fold action, they are not only anti-inflammatory, but they
are also pro-resolving of inflammation.
11, 12
Upon injury or infection, neutrophils in the blood
stream accumulate at the site of inflammation. The anti-inflammatory action of these distinct
lipid mediators decreases the infiltration of the neutrophils.
12
Pro-resolution of inflammation is
also enhanced by lipid mediators recruiting macrophages to engulf the apoptotic neutrophils and
these actions clear the site of inflammation non-phlogistically, meaning without heat or fever.
12

Nature Reviews | Immunology
Neutrophil
Monocyte
Endothelial cell
Neutrophil
infiltration
Monocyte
recruitment
Dual-acting
mediators
• Lipoxins
• Resolvins
• Protectins
Exudate
Capillary
Apoptotic
neutrophil
Uptake of apoptotic
neutrophils
Macrophage
Exit via
lymphatics
Exudate
Biological fluid that filters
from the circulatory system
into lesions or areas of
inflammation. Exudate is
characterized by a high content
of plasma proteins, cells and
cellular debris. Pus is an
example of an exudate found
in infected wounds and it
contains bacteria and high
concentrations of white blood
cells.
Prostaglandins
Cyclopentane-ring-containing
lipids derived from the
metabolism of arachidonic
acid by the action of
cyclooxygenases and
downstream synthase
enzymes. They have a diverse
range of biological activities
and a well-recognized role in
inflammation and pain.
cells and leukocytes
14
. This activity is enhanced during
inflammation but is also likely to provide a mechanism
for restoring homeostasis, because mucosal surfaces are
continuously exposed to microorganisms in vivo
29–31
. In
the mucosa, lipoxins are generated by neutrophils from
the 15-hydroxyeicosatetraenoic acid (15-HETE) precur-
sor, which is provided by mucosal epithelial cells
29
. The
blood vessels represent another main site where lipoxin
biosynthesis occurs in humans; biosynthesis involves
the initiation of arachidonic-acid oxygenation by
5-lipoxygenase in leukocytes and the release of the inter-
mediate leukotriene A
4
, which is converted by the lipoxin-
synthase activity of 12-lipoxygenase in platelets. This
biosynthesis pathway is exemplified by platelet–leukocyte
interactions in blood vessels or possibly in exudates that
form a nidus for transcellular biosynthesis.
In inflamed sites, neutrophils can interact with other
cells in their immediate vicinity, such as with other
leukocytes, platelets, endothelial cells, mucosal epithe-
lial cells and fibroblasts, and acquire the ability to pro-
duce lipoxins. More than 50% of the leukocyte-derived
epoxide intermediate leukotriene A
4
is released by cells
for processing by platelet 12-lipoxygenase or mucosal
15-lipoxygenase to produce lipoxins
32 34
. Lipoxins gen-
erated by cell–cell interactions and transcellular biosyn-
thesis stop neutrophil diapedesis and recruitment into
the tissues
29 31
(FIG. 1).
It is now clear that neutrophils change their phenotype
to produce different profiles of lipid mediators depend-
ing on the cells and substrates present in their local
environment
8,9
. For example, neutrophils in resolving
inflammatory exudates switch from the production of
leukotrienes to that of lipoxins and resolvins, whereas
neutrophils in the peripheral blood, on activation, generate
and release leukotriene B
4
as one of their main bioactive
products
8
. In this context, local prostaglandin E
2
and prosta-
glandin D
2
stimulate the processing of 15-lipoxygenase
mRNA in leukocytes to produce a functional enzyme for
lipoxin production
8
. Other cell types can acquire the abil-
ity to generate lipoxins when exposed to specific cytokines
or growth factors
32
, or in the case of macrophages, for
example, when they engulf apoptotic leukocytes
35
. These
findings are of relevance to pro-resolution mechanisms
because lipoxin A
4
generated by macrophages probably
contributes to the stimulation of their phagocytic activ-
ity
15,17
without elaborating pro-inflammatory mediators
— namely, the non-phlogistic process.
Pathogens can also contribute to the provision of
the necessary components for lipoxin biosynthesis.
Pseudomonas aeruginosa encodes the first identified
secretory lipoxygenase that converts host arachidonic
acid to 15-HETE for local lipoxin production
36
. High
levels of lipoxins, greater than those considered to be
physiologically standard, are produced by host cells
infected by Toxoplasma gondii, which encode their own
15-lipoxygenase
37,38
. Therefore, 15-lipoxygenase expressed
by pathogens may interact with endogenous biosynthetic
circuits of the host to generate local ‘stop signals’ at levels
that can divert the host immune defence.
Pro-resolution actions of lipoxins. Lipoxin A
4
and
lipoxin B
4
inhibit neutrophil entry into inflamed sites
and counter-regulate the main aspects of inflammation.
They act on many cell types including blood cells, neural
cells and stromal cells
39 42
(TABLE 1). Lipoxin A
4
regulates
leukocyte responses in vitro and trafficking in vivo by
activating its specific receptor, lipoxin A
4
receptor (ALX)
(FIG. 3). ALX is a G-protein-coupled receptor (GPCR) that
is expressed by leukocytes and has cell-type-specific
signalling pathways
40
. For example, in neutrophils,
lipoxin-A
4
–ALX interactions stop neutrophil migration,
whereas in monocytes lipoxin-A
4
–ALX interactions
stimulate monocyte chemotaxis and non-phlogistic
responses
41
. Unlike classic GPCRs for chemoattractants
that mobilize intracellular Ca
2+
to evoke chemotaxis,
lipoxins instead induce changes in the phosphorylation
of proteins of the cytoskeleton, resulting in cell arrest
42,43
.
In addition to these effects on the resolution of inflam-
mation, lipoxin A
4
reduces organ fibrosis, acts directly
on both vascular and smooth muscle (for reviews, see
REFS 14,40) and has direct action in reducing pain
44
.
Aspirin impinges the endogenous lipoxin-generating
system during cell–cell interactions. Inhibition of pros-
taglandin biosynthesis by aspirin is a well-appreciated
mechanism in its anti-thrombotic and anti-inflammatory
effect
45
. Of interest, aspirin triggers the endogenous
formation of carbon-15 epimeric lipoxins, namely
aspirin-triggered lipoxins (ATLs)
46
. Cells that express
cyclooxygenase-2 (COX2), including vascular endothe-
lial cells, epithelial cells, macrophages and neutrophils,
are involved in ATL production. Acetylation of COX2 by
aspirin blocks its ability to biosynthesize prostaglandins
Figure 2 | Dual anti-inflammatory and pro-resolution actions of specific lipoxins,
resolvins and protectins. The key histological feature in the resolution of inflammation
is the loss of neutrophils from the local inflamed sites. This is a programmed process
that is actively regulated at multiple levels: by reducing neutrophil infiltration into the
exudate, increasing monocyte recruitment to the exudate, stimulating macrophage
uptake of apoptotic neutrophils, and promoting phagocyte exit from the exudate via
the lymphatics.
REVIEWS
352 | MAY 2008 | VOLUME 8 www.nature.com/reviews/immunol
6
By this action, lipid mediators produced by 15-LOX-1 not only stop the inflammation, but also
help to clear the site of inflammation.

1.3 DHA-Derived Lipid Mediators
DHA is one of the most important PUFAs because it gets biosynthesized into resolvins,
maresins, and protectins. A few of the notable lipid mediators that are derived from DHA are
resolvins D1, and D2, as well as maresin 1 and neuroprotectin 1. For example, DHA is
transformed into neuroprotectin D1 (NPD1) via an enzymatic process, shown in Scheme 1.1.
1

NPD1 is critical to healthy brain function because it attenuates brain injuries and disorders, such
as Alzheimer’s disease and stroke.
13, 14
Although NPD1 is derived from DHA, it was found that
NPD1 is much more potent biologically than DHA.
11

Scheme 1.1- Biosynthesis of neuroprotectin D1 (NPD1) from DHA.
1

Scheme 1.1 outlines the synthesis of NPD1 from DHA, starting with the stereospecific
peroxidation at carbon-17 by 15-LOX using oxygen, producing 17S-Hydroperoxy-DHA (17S-
HpDHA).
1
There are two pathways that the 17S-HpDHA can then go through, one possibility is
7
that it is reacted with a peroxidase enzyme, then the peroxide gets reduced to the alcohol to yield
the 17S-Hydroxy-DHA (17S-HDHA).
1, 11
17S-HDHA is also an intermediate toward the
biosynthesis of RvD1 and RvD2. The other pathway that 17S-HpDHA undergoes is an
enzymatic epoxidation to produce the 16S, 17S-Epoxide shown in Scheme 1.1. The epoxide can
then be hydrolyzed enzymatically as well as stereoselectively oxidized at carbon-10, to yield the
10R, 17S-diHydroxyDHA (NPD1). The biosynthesis of NPD1 shown in Scheme 1.1 is just one
of the pathways in which DHA is converted into a lipid mediator. However, some known
metabolic pathways of DHA also begin with 15-LOX and some go through the same
intermediates.
One DHA metabolic pathway which does not begin with 15-LOX is within the
production of D-series aspirin-triggered resolvins (AT-RvD). The D-series resolvins are so-
named because they are derived from DHA, and thus have a docosahexaenoic acid backbone.
The term “aspirin-triggered” comes from the fact that these lipid mediators are produced when
the enzyme cyclooxygenase-2 (COX-2) is exposed to aspirin.
1
The enzyme cytochrome P450
can also have the same effect as COX-2 in the first step of this pathway.
1
The enzymatic
pathway detailing how DHA is metabolized into AT-RvD1 through AT-RvD4 is shown in
Scheme 1.2.
1

In the first step of the pathway the enzyme P450 or COX-2 and aspirin, can use oxygen to
stereoselectively peroxidate the 17-position of DHA to produce 17R-HydroperoxyDHA (17R-
HpDHA). Peroxidase enzyme can then reduce the peroxide on 17R-HpDHA to the 17R-
HydroxyDHA (17R-HDHA). From here, the 17-hydroxy intermediate reacts with 5-
lipoxygenase (5-LO) in the presence of oxygen to produce two different epoxides. 5-LO
epoxidizes the 7,8-position stereoselectively, to result in 7S,8S-Epoxide, which gets hydrolyzed
8
to the lipid mediator products AT-RvD1 and AT-RvD2. However, 5-LO also epoxidizes the 4,5-
position of 17R-HDHA, and depending on the position of the hydrolysis of the 4,5-Epoxide from
the hydrolase enzyme, either AT-RvD3 or AT-RvD4 is produced.

Scheme 1.2- Aspirin-triggered resolvin D-series compounds biosynthesized from DHA.
1

1.4 AA-Derived Lipid Mediators
Arachidonic acid (AA) is an omega-6 fatty acid that is unique because unlike omega-3
fatty acids it can be converted into both pro- and anti-inflammatory lipid mediators. Figure 1.1
shows an overview of some of the possible pathways that AA can go through and Figure 1.3
details the structures of the lipid mediators produced.
1, 4, 15
AA interacts with several enzymes to
5930 dx.doi.org/10.1021/cr100396c |Chem. Rev. 2011, 111,5922–5943
Chemical Reviews
REVIEW
forms 28 that is similarly converted to RvD3, RvD4, and
RvD6.
Aspirin-triggered 17R D-series resolvins were also identified
(Figure 8).
4,79
Initial oxygenation at C-17 to form 17R-HpDHA
(30)takes place in the presence of aspirin via acetylated
COX-2 or via a P450 pathway. Subsequent enzymatic
transformations lead to AT-RvD1 (33), AT-RvD2 (34), and
other AT-resolvins.
3.2. Resolvin D1
Resolvin D1 (RvD1) is biosynthesized in situ from DHA by
two sequential oxygenations at the carbon 7 and 17 positions
Figure 7. Pathways and enzymes in the biosynthesis of D-series resolvins.
Figure 8. Pathways and enzymes involved in the biosynthesis of aspirin-triggered D-series resolvins.
9
produce such structurally diverse lipid mediators. The stereochemistry of each stereogenic
center on the arachidonic acid metabolites is an indication of which enzyme metabolized the
fatty acid. In the case of lipoxin A
4
compounds (Figure 1.3, shown in green), the
stereochemistry at carbon-15 is dependent on the first enzyme to interact with AA. 15-LOX-1
gives the S-configuration at C-15, and later produces LXA
4
, whereas COX-2 in the presence of
aspirin yields the R-configuration at C-15 and forms the aspirin-triggered-LXA
4
. However, both
R- and S-configuration intermediates can then interact with 5-LO to result in the same 5S,6S-cis-
diol moiety near their carboxylic acid part of the chain.
15

Figure 1.3- Arachidonic acid and the lipid mediators it is metabolized into.

Leukotrienes are lipid mediators that have three conjugated double bonds in their
structure. In order to form the pro-inflammatory leukotrienes (Figure 1.3, shown in red), AA is
COOH
Arachidonic Acid (AA)
COOH
OH HO
OH
Lipoxin A
4
(LXA
4
)
COOH
OH HO
OH
15-epi-Lipoxin A
4
(AT-LXA
4
)
OH
COOH
OH
Leukotriene B
4
(LTB
4
)
COOH
OH
S
NH
O
N
H
O
OH
O
H
2
N
O
OH
Leukotriene C
4
(LTC
4
)
O
HO
OH
COOH
Prostaglandin E
2
(PGE
2
)
O
HO
OH
COOH
Prostaglandin D
2
(PGD
2
)
10
stereoselectively oxygenated at the 5-position by 5-LO to produce leukotriene A
4
.
15
Leukotriene
A
4
is a key intermediate because it can become the anti-inflammatory LXA
4
, as well as the pro-
inflammatory leukotrienes B
4
and C
4
(LTB
4
and LTC
4
). LTB
4
is responsible for recruiting
neutrophils as well as other types of cells to the site of inflammation, and thus is critical to the
onset and early stages of the inflammatory response.
15

LTC
4
is a cysteinyl leukotriene, meaning
it has cysteine in its structure, and it binds to receptors and causes muscle contractions in specific
tissues.
15

Another class of lipid mediators that are derived from AA is prostaglandins, which are
abundant all over the body in most tissues and cells. Figure 1.3 highlights two of the several
prostaglandins that AA is metabolized into, prostaglandins D
2
and E
2
(PGD
2
and PGE
2
shown in
pink). Prostaglandins are synthesized via AA binding to COX-1 or COX-2 and undergoing
several radical reactions to produce the intermediate prostaglandin H
2
(PGH
2
). PGH
2
can bind to
specfic synthesases to produce the bioactive PGD
2
or PGE
2
. Prostaglandin D
2
has many diverse
and important purposes biologically, from causing uterine contractions to inducing fever.
Conversely, prostaglandin E
2
is just as vital to biological function because it causes a reduction
in body temperature as well as vasodilation. Polyunsaturated fatty acids have a significant role
in many biological processes and the slightest perturbance in their delicate balance can cause
severe problems to health and normal function.

1.5 Conclusion
Polyunsaturated fatty acids and lipid mediators not only play a critical role in
inflammation, but also have been found to be of great significance in many diverse biological
11
processes. Although several inflammatory pathways and metabolites have been elucidated, the
link between inflammation and certain diseases is still unclear. Lipid mediators are critical to
cellular signaling pathways and therefore further research is required to discover novel lipid
metabolites and their structure-activity-relationships within the biological system. Furthermore,
lipid mediators hold great potential not only in the field of inflammatory diseases but also for
other therapeutics as well. Overall, future research elucidating novel signaling pathways and
lipid metabolites could help determine the connection between inflammation, lipoxygenating
enzymes, and cancer.

12
1.6 Chapter 1 References

1
Serhan, C. N.; Petasis, N. A. Chem. Rev. 2011, 111, 5922.

2
Burr, G. O.; Burr, M. M. J. Biol. Chem. 1929, 82, 345.

3
Dyerberg, J.; Bang, H. O.; Hjorne, N. Am. J. Clin. Nut. 1975, 28, 958.

4
Finaldi, A.-M.; Petasis, N. A. 243
rd
ACS National Meeting and Exposition, San Diego, CA,
March 2012.

5
Uderhardt, S.; Kronke, G. J. Mol. Med. 2012, 90, 1247.

6
Jiang, W. G.; Watkins, G.; Douglas-Jones, A.; Mansel, R. E. Prostaglandins Leukot. Essent.
Fatty Acids 2006, 74, 235.

7
Funk, C. D. Arterioscler. Thromb. Vasc. Biol. 2006, 26, 1204.

8
Kelavkar, U. P.; Nixon, J. B.; Cohen, C.; Dillehay, D.; Eling, T. E.; Badr, K. R. Carcinogenesis
2001, 22, 1765.

9
Tang, S.; Bhatia, B.; Maldonado, C. J.; Yang, P.; Newman, R. A.; Liu, J.; Chandra, D.; Traag,
J.; Klein, R. D.; Fischer, S. M.; Chopra, D.; Shen, J.; Zhau, H. E.; Chung, L. W. K.; Tang, D. G.
J. Biol. Chem. 2001, 277, 16189.

10
Levy, B. D.; Clish, C. B.; Schmidt, B.; Gronert, K.; Serhan, C. N. Nature Immun. 2001, 2, 612.

11
Serhan, C. N.; Yacoubian, S.; Yang, R. Annu. Rev. Mech. Dis. 2008, 3, 279.

12
Serhan, C. N.; Chiang, N.; Van Dyke, T. E. Nature Immun. 2008, 8, 349.

13
Zhao, Y.; Calon, F.; Julien, C.; Winkler, J. W.; Petasis, N. A.; Lukiw, W. J.; Bazan, N. G.
PLoS One 2011, 6, e15816.

14
Bazan, N. G. J. Lip. Res. 2009, 50, S400.

15
Petasis, N. A.; Akritopoulou-Zanze, I.; Fokin, V. V.; Bernasconi, G.; Keledjian, R.; Yang, R.;
Uddin, J.; Nagulapalli, K. C.; Serhan, C. N. Prostaglandins Leukot. Essent. Fatty Acids 2005, 73,
301.

13
Chapter 2. A Docosahexaenoic Acid Analog as an Imaging Probe
for Lipid Mediator Pathways
2.1 Introduction
Docosahexaenoic acid (DHA) is an omega-3 fatty acid that is vital to brain and eye
development and function. DHA constitutes the majority of polyunsaturated fatty acids (PUFAs)
in the human brain and retina, 40% and 60% respectively, and 50% of neuronal membrane
weight.
1
Oxygenated metabolites of DHA (such as resolvins, lipoxins, protectins, and maresins)
have many beneficial anti-inflammatory and cyto-protective properties essential to human health
(Scheme 2.1).
2, 3, 4, 5, 6
In 2003, it was discovered that DHA in the brain is metabolized into
neuroprotectin D1 (NPD1), so named because of its protective effects on the brain during
oxidative stress.
7, 8

Scheme 2.1- DHA and a few of the lipid mediators it is metabolized into (resolvins D1 and D2,
neuroprotectin D1, and maresin R1).
9, 10

Although several metabolites of DHA have been widely studied, new metabolites are
being discovered presently, and the mechanism of distribution and trafficking of lipid mediators
is not completely understood. Also, understanding what proteins new lipid mediators are
signaling to could uncover new pathways or new targets that have not been investigated before.
OH
HO
COOH
HO
COOH
HO
OH
HO
OH
OH
COOH
OH COOH
OH
COOH
RvD1
RvD2 NPD1
MaR1
DHA
14
By tagging DHA we can use UV or fluorescence to identify new DHA derived metabolites.
However, for such a molecule to be effective, there are many requirements that need to be met.
i.) The molecule must contain a “tag” on the omega (terminal) end of the DHA.
The tag cannot be attached to the alpha (carboxylic acid) end of the DHA,
because this would interfere with the reactivity of the carboxylic acid. Also,
the tag itself must be small and not interfere with DHA’s metabolism or uptake
either.
ii.) Ideally, the molecule should fluoresce only when we desire to detect it.
iii.) The chemical reporter that will react with the tag should give a specific
wavelenth emission, which would allow for quantification of low-level
quantities of metabolites via a fluorometer.
iv.) The tagged metabolites should have a different molecular weight and UV
absorbance compared with their untagged counterparts, which would allow for
analysis and identification of the tagged metabolites via LC-MS/MS, without
the confusion of the untagged metabolites interfering with the data. This
should also allow for the quantification of the ratios of different metabolites
present, which is highly important because the disturbance of the delicate
balance of lipid mediators can indicate a stressed or diseased environment, as
discussed in Chapter 1.

These criteria can be met through the use of “click chemistry”, involving a Huisgen [3+2]
cycloaddition reaction catalyzed by copper(I).
11,

12
A representative example of a “click”
reaction is shown above in Scheme 2.2. Generally, a terminal alkyne and an azide are reacted
15
together in the presence of copper(I) to yield a 1,2,3-triazole. Although click chemistry has
found several significant diverse applications, more recently it has been utilized as a method for
chemical reporting and molecular imaging. For example, an alkyne “tag” is incorporated into a
molecule of interest (the probe), and an azide (reporter) contains a non-fluorescent fluorophore.
After the tagged molecule has been metabolized, the fluorophore is “clicked” to the alkyne
metabolite, and product results in a conjugated ring system that emits fluorescence. Of course
the situation can also be reversed, wherein the azide is the probe and the alkyne is the reporter,
but the “click” can still result in fluorescence.

Scheme 2.2- A general example of a “click” reaction.

Coumarin azide analogs (shown in Scheme 2.3) have picked up some interest in the field
of click chemistry specifically as non-fluorescent fluorophores that fluoresce upon formation of
the 1,2,3-triazole ring.
13
A general example of an alkyne probe reacting with a coumarin azide
(the non-fluorescent fluorophore) to yield a fluorescent 1,2,3-triazole is shown in Scheme 2.3.
13

We were interested in using a coumarin azide as a reporter due to their ease of synthesis and
well-established practicality.
R
1
N N N
R
2
Cu(I)
+
N
N
N
R
1
R
2
16

Scheme 2.3- Click chemistry that results in a fluorescent product.
13

Probing undiscovered pathways is helpful to elucidating disease pathogenesis and
treatment. The more complex the model system used, the more accurate the results will be in
relation to a human patient. While cell studies are a useful tool, cells do not hold the complexity
and signaling pathways between different types of tissues and organs. A mouse model could give
a more complete picture of a full signaling pathway, and therefore there is a great amount of use
for click chemistry in living animals. Recently, Bertozzi and co-workers have done some
groundbreaking work in the area of chemical reporters via copper-free click chemistry.
14,

15

Through the use of strained cyclooctyne and difluorocyclooctyne in place of the unstrained
terminal alkyne, it is possible to carry out the click reaction in a timely manner without the use of
cytotoxic copper. Bertozzi and co-workers also demonstrated the first copper-free click reaction
in living animals, thereby broadening the concept of click chemistry and how it can be used
(Figure 2.1).
15

R
1
N
N
N
Cu(I)
+
N
N
N
R
1
O O OH
O O OH
Alkyne
(probe)
Non-fluorescent
fluorophore
azide
(reporter)
Clicked fluorescent
product
17

Figure 2.1- Bertozzi’s copper-free click chemistry in living animals.
15

Although Bertozzi’s copper-free click chemistry in living animals did not include any
fluorescent probes, much of her previous work has utilized fluorescence, and her strained
cyclooctyne system could be adapted for fluorescence in animals as well.
14, 15
Achieving click
chemistry in a living animal proves that this reaction could possibly answer many more questions
regarding the mechanisms of metabolism and diseases. Such work heralds several possibilities
for future applications of click chemistry relating to disease diagnosis, cancer treatment, as well
as elucidation of unknown molecular pathways.
Given the promise of click chemistry, we investigated its application to DHA in order to
enhance the probing of DHA metabolism.

18
2.2 Results and Discussion
2.2.1 Design of an ω-Alkynyl-DHA Derivative
The structure of DHA is deceptively simple, and yet very difficult to synthesize in high
yield with complete stereocontrol over the six all-cis double bonds. While it is easy to alter the
structure of commercially available DHA at the carboxylic (alpha) end, it is much more
challenging to instill a new functional group towards the omega end of the molecule. In order to
incorporate a terminal alkyne onto the omega end of DHA, the molecule must be fully
synthesized. It is important to note that DHA is a linear molecule, and thus requires a mainly
linear synthesis. Although pieces can be synthesized individually, they can only be added one at
a time, linearly. Also, the synthesis must retain the cis configuration about all of the double
bonds, which can easily be isomerized to the more stable trans isomer. Great care must be given
to ensure that the double bonds and terminal alkyne are not oxidized or exposed to harsh
conditions, which could result in polymerization or degradation of the entire product.
Current scientific literature has minimal amounts of publications regarding the total
synthesis of DHA derivatives when compared to other types of molecules. The main reason for
this is that the total synthesis of DHA is arduous and DHA itself is commercially available and
also easily biosynthesized from proteins and cells. Previous syntheses of DHA and its
derivatives utilized Wittig reactions to install the six cis double bonds.
16
The Wittig reaction
requires great care, very specific conditions, and often results in a mixure of cis and trans
products that are problematic to separate. Alternatively, Lindlar hydrogenations can yield a cis
alkene exclusively, but suffer from also producing an over-hydrogenated side product.

19
The ω-alkynyl-DHA probe (2.1) and its retrosynthesis are shown in Scheme 2.4. The
trimethylsilyl (TMS) protected bromide (2.2) is utilized three times in the overall synthesis, and
is derived from the commercially available TMS-propargyl bromide. Due to this, a large
quantity of 2.2 could be synthesized easily and used as needed throughout the synthesis. All of
the cis double bonds were produced via selective Lindlar hydrogenations, which provided a
“handle” for reactivity as well as a way to assemble the terminal alkyne. Consecutively building
off of a terminal alkyne also allowed for utilization of copper-mediated coupling reactions. My
synthetic strategy aided in my completion of the first total synthesis of an ω-alkynyl-DHA
imaging probe.

Scheme 2.4- A retrosynthetic analysis of the alkynyl-DHA derivative target.

2.2.2 First Total Synthesis of an ω-Alkynyl-DHA Derivative as an Imaging Probe
The total synthesis of an ω-alkynyl-DHA derivative (2.1) is shown in Schemes 2.5 and
2.6. Beginning with commercially available TMS-propargyl bromide and propargyl alcohol, a
copper-mediated coupling reaction was used to produce a skipped di-yne alcohol (2.3). The
reaction was run with CuI, NaI, and K
2
CO
3
in N,N’-dimethylformamide (DMF), at room
temperature overnight to give 2.3 in good yield. The alcohol was then converted into the
COOMe
TMS
Br
TMS
Br
2.2
COO
-
Na
+
2.1
20
bromide (2.2) by treatment with triphenylphosphine and n-bromosuccinimide (NBS) in
dichloromethane at 0
o
C, in modest yield. The bromide 2.2 was an essential building block for
the majority of the total synthesis. The copper-mediated coupling was then repeated with the
same conditions to couple 4-pentynoic acid methyl ester with 2.2 to attain the TMS-protected
skipped tri-yne (2.4).

Scheme 2.5- The first half of the total synthesis of ω-alkynyl-DHA (2.1), a potential imaging
probe for lipid mediator pathways.

Importantly, the skipped tri-yne (2.4) needed to be reduced selectively to two skipped cis
double bonds, leaving the TMS-protected alkyne intact, which would allow for a deprotection of
the terminal alkyne and another copper-mediated coupling reaction. The selective hydrogenation
was achieved via a Lindlar catalyst hydrogenation, wherein the TMS group protects the terminal
alkyne from hydrogenation. Lindlar catalyst was added to a solution of 2.4 in ethyl acetate at
room temperature, and a small amount of quinoline was added in order to deactivate the catalyst
and enhance the selectivity of the reaction by preventing over-hyrdogenation to the alkane side
product. A hydrogen gas balloon was added to the set up, and TLC was used to monitor the
Br
TMS a TMS
OH
b TMS
Br
a
COOMe
2.3 2.2
COOMe
TMS c, d
COOMe
2.4
2.5
Reagents and condtions: (a) K
2
CO
3
, CuI, NaI, DMF, rt, overnight, 86% and 51% yield (b) PPH
3
, NBS, DCM, 0
o
C to rt,
30 min., 37% yield (c) Lindlar's Cat., H
2
, quinoline, EtOAc, rt (d) Na
2
CO
3
, MeOH, rt, overnight 26% (over two steps)
21
reaction closely. Production of the singly-hydrogenated, doubly-hydrogenated (product), and
over-hydrogenated side product were carefully visualized on the TLC plate. Even with careful
monitoring and several precautions taken, such as limited catalyst and adding extra quinoline,
prevention of over-hydrogenation was difficult and this reaction almost always resulted in low or
modest yields. However, the advantages to using a selective Lindlar catalyst hydrogenation is
that the cis isomer is produced exclusively, and also the over-hydrogenated side product is easily
separable from the desired product via column chromatography. Another advantage to this
method is that the crude product does not need to be purified prior to removal of the TMS
functional group. The protected product was stirred under mild conditions with dry methanol
and sodium carbonate to yield 2.5.
With the acetylenic bis-cis-alkene (2.5) in hand, the following step was another copper-
mediated coupling with the essential building block 2.2 (Scheme 2.6). The purified product was
the TMS-protected skipped tri-yne with two skipped cis-alkenes (2.6). Another selective
hydrogenation using Lindlar’s catalyst was employed to reduce the two internal alkynes and
leave the TMS-protected terminal alkyne untouched. Compound 2.7 was synthesized
successfully, after removal of the TMS group from the crude product, using the exact same
conditions as previously.

22

Scheme 2.6- The second half of the total synthesis of the title compound (2.1).

Reaching close to the end of the synthesis, one last copper-mediated coupling reaction
with 2.2 was necessary to install all of the functionality necessary to finish the synthesis. Upon
completion of purified 2.8, the same selective hydrogenation and deprotection methods were
utilized to yield the methyl ester analog (2.9) of the title compound 2.1. The methyl ester (2.9)
was purified via HPLC, and the pure product was reacted with sodium hydroxide to yield the
sodium salt 2.1, which was also purified by HPLC. The title compound (2.1) was successfully
synthesized completely, with high purity and selectivity.

TMS
COOMe
2.2, a
2.5
COOMe
TMS
COOMe
2.7
2.6
COOMe
2.8
Reagents and condtions: (a) K
2
CO
3
, CuI, NaI, DMF, rt, overnight, 56% and 26% yield (b) Lindlar's Cat., H
2
, quinoline,
EtOAc, rt (c) Na
2
CO
3
, MeOH, rt, overnight 54% and 18% (over two steps) (d) NaOH, H
2
O, EtOH, 0
o
C, 48 hrs., 15%
b, c
2.2, a
COOMe
2.9
b, c
d
COO
-
Na
+
2.1
23
2.2.3 Lipoxygenation of an ω-Alkynyl-DHA Derivative with 15-Lipoxygenase
In light of the recent advancements of click chemistry, we sought to design a DHA
molecule that could probe pathways, fluoresce where it is being metabolized, and possibly be
used in animal studies. Figure 2.2 shows the target molecule the general approach to how we
envisaged utilizing it. Ideally, our ω-alkynyl-DHA could be “clicked” in vivo to yield a triazole
that would be easily detected and quantified. Our enzyme of interest was 15-lipoxygenase (15-
LO) (described in more detail in chapter 1), due to its ability to metabolize DHA into highly
potent anti-inflammatory agents such as resolvins and neuroprotectin 1. It was necessary to
determine if the ω-alkynyl-DHA derivative would be metabolized in the same way as standard
DHA. 15-LO metabolizes DHA mainly into the 17-hydroxylated product, therefore our goal was
for the ω-alkynyl-DHA to also produce the 17-hydroxylated product when reacted with 15-LO in
vitro. If the terminal alkyne tag interfered in the metabolism or hindered the activity of the
molecule in any way, the effectiveness of the molecule would be nullified.

Figure 2.2- A novel DHA analog as an imaging probe for lipid mediator pathways.
10
The blue
bead and star represent a fluorophore.

a
b
COOH COOH
N
N
N
COOH
OH
17
N
3
COOH
N
N
N
OH
17
15-LO O
2
& other metabolites
COOH
COOMe
Br
TMS
Br
TMS
OH
TMS
Br
TMS
COOMe
a
TMS
COOMe
COOMe
OH
c,d
Br
TMS
a
COOMe
TMS
c,d
COOMe
Br
TMS
COOMe
TMS
c,d,e
COOH
Reagents & Conditions: (a) K
2
CO
3
, CuI, NaI, 56-70%; (b) PPh
3
, NBS, DCM, 30-40%;
(c) Lindlar's Cat., H
2
, quinoline, EtOAc; (d) Na
2
CO
3
, MeOH; (e) NaOH, H
2
O, MeOH; (f)
DPPA, DIAD, PPh
3
, THF.
COOMe
a
a OH
COOMe
Cl OH
c
OH
COOMe
N
3
COOH
f,e
!-Alkynyl DHA
!-Azido DHA
Br
TMS
24
Importantly, the activity of the molecule needed to be tested enzymatically before it could
be tested in vivo. Not only is an enzymatic assay cheaper than cell studies, but it also gives the
possibility to rule-out unsuccessful probe candidates. Also, an enzyme study gives clean
metabolic products, which are much easier to extract and interpret rather than cell studies, and
these metabolites can dictate how to redesign the molecule if necessary. By choosing an enzyme
that is known to have a high level of activity of metabolizing DHA into vital anti-inflammatory
agents, it is possible to learn if the acetylenic tag would affect the metabolism of the molecule.
15-Lipoxygenase (15-LO) is an enzyme that plays a critical role in the metabolism of DHA into
anti-inflammatory resolvins. 15-LO uses oxygen to form peroxides at the double bonds on
DHA, and then the peroxide bond can be reduced via sodium borohydride to the hydroxyl
product. In vivo this reduction to the hydroxylated product takes place through a different
method, but with only the enzyme present in vitro, the reduction should be performed manually.
Prodominantly, 15-LO produces the mono-hydroxyl product 17-hydroxydocosahexaenoic acid
(17-HDHA), but also minor amounts other hydroxylated products including the di-hydroxy
metabolites (10,17-HDHA and 7,17-HDHA). Using work recently published by Dangi, et. al. as
a model reaction between standard DHA and 15-LO, I was able reproduce their results and
biosynthesize the 17-HDHA, our metabolite of interest.
17
The product was analyzed via LC-
MS/MS, and the fragmentation confirmed the presence of the major product 17-HDHA (See
Scheme 2.7).
Once the model reaction was confirmed successful, the same reaction could be tested
with the ω-alkynyl-DHA derivative to investigate whether or not the metabolism would be
affected due to the variation in structure. The most critical test for validation of the probe’s
possible success is that it is metabolized mainly into the 17-hydroxy-ω-alkynyl-DHA, even if the
25
minor (di-hydroxy) products predominated, then the probe would be unsuccessful and need to be
completely redesigned. As shown in Scheme 2.7, it was found that the acetylenic tag did not
hinder the metabolism of the DHA probe in any way. The major product of the enzymatic
reaction of 15-LO with ω-alkynyl-DHA was still the 17-hydroxy metabolite. The product was
observed via LC-MS/MS and the fragmentation pattern for the alkyne-tagged DHA was the same
as the model reaction with the standard DHA. The mass spectrometry spectrum of 2.11 is shown
in Scheme 2.7 and in the appendix of this work. The positive lipidomic analysis indicated that
the ω-alkynyl-DHA would be an acceptable probe to use for further studies, including cell-based
assays.

26

Scheme 2.7- Lipoxygenation reaction with 15-lipoxygenase (15-LO) and standard DHA or ω-
alkynyl-DHA.

COO
-
2.1
COO
-
DHA
a) 15-LO
sodium borate buffer
b) NaBH
4
COO
-
OH
17-HDHA
245
273
2.10
COO
-
OH
245
273
2.11
Exact mass: 343.2279
UV: 237 nM
Exact mass: 353.2122
UV: 237 nM
a) 15-LO
sodium borate buffer
b) NaBH
4
27
2.3 Conclusion
In conclusion, the first total synthesis of a novel DHA-derivative for use as an imaging
probe of lipid mediator pathways has been achieved. Selective Lindlar catalyst hydrogentations,
as well as copper-mediated coupling reactions, were utilized to execute the synthesis in twelve
overall steps. The highly purified product (2.1) was subjected to enzymatic studies, to ensure the
akynyl tag would not affect the metabolism of the molecule. Upon discovering that the
metabolism of the ω-alkynyl-DHA derivative was identical to that of standard DHA, the new
analog has been identified as a possible probe for both cell and animal assays. Further biological
studies are currently underway to utilize this interesting probe to elucidate pathways of lipid
mediators via click chemistry.

2.4 Experimental
All reactions, unless otherwise noted, were carried out with commercially available
starting materials and solvents, which were used without further purification. The majority of
the chemicals were purchased from Sigma Aldrich, GFS chemicals, TCI America, and VWR
International. All glassware and reaction vessels used were standard and commercially
available. Reactions performed under “inert atmosphere” implies the reaction was run in a round
bottom flask affixed with a three-way adapter and an argon balloon, or the flask was affixed with
a septum which was punctured with a needle affixed to an argon balloon.
1
H and
13
C NMR
spectra were collected using a Varian Mercury 400, Varian 400-MR, Varian 500, or Varian 600
NMR spectrometers, using residual
1
H and
13
C peaks of deuterated solvents as internal standards.
Manual flash column chromatography was run with silica gel purchased from Sorbent
28
Technologies (60 Å, 40-63 µm). A Biotage Isolera One was used for automated flash column
chromatography, with standard Biotage silica cartridges (60 Å) for purification. LC-MS analysis
was performed on an Agilent Technologies 1200 infinity series instrument. LC-MS/MS analysis
was performed on a Finnigan LCQ Deca XP Max mass spectrometer with a Finnigan Surveyor
PDA 158 Plus detector.

Procedures

6-(trimethylsilyl)hexa-2,5-diyn-1-ol (2.3). K
2
CO
3
(8.133 g, 58.84 mmol), NaI (8.820 g, 58.84
mmol), and CuI (11.324 g, 58.84 mmol) were suspended in DMF (50 mL) under inert and
anhydrous atmosphere. Propargyl alcohol (2.82 mL, 39.21 mmol) was added via syringe to the
slurry, and allowed to stir for 30 minutes at room temperature. Next, TMS-propargyl bromide
(5.18 mL, 36.65 mmol) was added dropwise, and the reaction was stirred overnight at room
temperature. The reaction was quenched with saturated aqueous ammonium chloride and then
extracted with diethyl ether (3 x 50mL). The combined organic layers were dried over
magnesium sulfate, filtered, and then concentrated in vacuo. The crude product was purified by
silica gel column chromatography run at 15% ethyl acetate:hexanes, to yield a colorless oil (5.26
g, 86%).
1
H NMR (400 MHz, Chloroform-d) δ
H
4.27 (t, J = 2.2 Hz, 2H), 3.26 (t, J = 2.2 Hz,
2H), 0.16 (s, 9H).
13
C NMR (151 MHz, Chloroform-d) δ 99.28, 85.47, 79.70, 78.82, 51.31,
10.96, -0.13.

TMS
OH
29

(6-bromohexa-1,4-diyn-1-yl)trimethylsilane (2.2). To a solution of 2.3 (3.695 g, 22.22 mmol)
in anhydrous DCM (40 mL), was cannulated PPh
3
(6.411 g, 24.44 mmol) in 5 mL of anhydrous
DCM at 0
o
C. N-Bromosuccinimide (4.350 g, 24.44 mmol) was quickly added in, and the
reaction was stirred at 0
o
C for 30 minutes. The reaction was quenched with saturated aqueous
sodium bicarbonate, extracted with diethyl ether (3 x 50 mL), and then washed with brine. The
combined organic layers were dried over magnesium sulfate, filtered, and then concentrated in
vacuo. The crude product was purified via silica gel column chromatography eluted with a
gradient of pure pentanes to 5% diethyl ether:pentanes. The purified oil (1.865 g, 37%) was
afforded as a slightly yellow oil.
1
H NMR (400 MHz, Chloroform-d) δ
H
2.65 (t, J = 7.2 Hz, 2H),
2.37 (t, J = 7.3 Hz, 2H), 0.24 (s, 9H).
13
C NMR (151 MHz, Chloroform-d) δ 98.72, 85.74, 80.86,
75.73, 14.71, 11.18, -0.16.

methyl 11-(trimethylsilyl)undeca-4,7,10-triynoate (2.4). Compound 2.4 was synthesized
similarly to compound 2.3, in 51% yield.
1
H NMR (500 MHz, Chloroform-d) δ
H
3.70 (s, 3H),
3.20 (t, J = 2.4 Hz, 2H), 3.12 (p, J = 2.3 Hz, 2H), 2.55-2.46 (m, 4H), 0.16 (s, 9H).
13
C NMR
TMS
Br
COOMe
TMS
30
(126 MHz, Chloroform-d) δ 172.41, 99.79, 85.13, 78.70, 74.99, 74.63, 73.87, 51.76, 33.35,
14.63, 10.90, 9.81, -0.11.

(4Z,7Z)-methyl undeca-4,7-dien-10-ynoate (2.5). To a solution of 2.4 (0.210 g, 0.81 mmol) in
ethyl acetate (10 mL) was added Lindlar catalyst (~ 50 mg) and 1 drop of quinoline. The
reaction was stirred under a hydrogen balloon and monitored closely by TLC until the doubly
hydrogenated compound was visualized. The reaction mixture was filtered through celite and
then concentrated in vacuo. To the crude, neat TMS-protected compound was added 10 mL of
HPLC grade methanol and 0.200 g (1.89 mmol) sodium carbonate, and the reaction mixture was
stirred at room temperature overnight. The reaction was filtered, concentrated in vacuo, and then
purified via column chromatography. The silica gel column was run at 4% diethyl
ether:pentanes, to afford 0.100g (0.52 mmol, 26% yield over two steps) of purified product 2.5 as
a colorless oil.
1
H NMR (500 MHz, Chloroform-d) δ
H
5.50-5.37 (m, 4H), 3.68 (s, 3H), 3.02 (t, J
= 0.5 Hz, 1H), 2.97 (ddt, J = 5.4, 2.9, 0.7 Hz, 2H), 2.83 (tdd, J = 5.5, 2.4, 1.4 Hz, 2H), 2.36 (d, J
= 1.1 Hz, 1H), 2.19 (td, J = 7.0, 2.7 Hz, 1H), 2.10-2.04 (m, 1H), 1.98 (t, J = 2.7 Hz, 1H).

(4Z,7Z)-methyl 17-(trimethylsilyl)heptadeca-4,7-dien-10,13,16-triynoate (2.6). Compound
2.6 was synthesized similarly to compound 2.3, in 56% yield.
1
H NMR (500 MHz, Chloroform-
COOMe
COOMe
TMS
31
d) δ
H
5.48-5.37 (m, 4H), 3.68 (s, 3H), 3.21 (t, J = 2.4 Hz, 2H), 3.14 (p, J = 2.4 Hz, 2H), 2.98-
2.92 (m, 2H), 2.82 (ddt, J = 5.5, 4.1, 2.6 Hz, 2H), 2.39-2.37 (m, 4H), 0.16 (s, 9H).
13
C NMR
(126 MHz, Chloroform-d) δ 173.50, 129.44, 128.69, 128.23, 124.72, 99.85, 85.11, 78.73, 75.12,
73.75, 68.13, 51.58, 33.97, 25.46, 22.65, 17.16, 10.92, 9.86, -0.10.

(4Z,7Z,10Z,13Z)-methyl heptadeca-4,7,10,13-tetraen-16-ynoate (2.7). Compound 2.7 was
synthesized similarly to compound 2.4, and collected as light yellow oil in 54% yield over two
steps (43 mg).
1
H NMR (600 MHz, Chloroform-d) δ
H
5.53-5.44 (m, 2H), 5.39 (tddd, J = 11.1,
9.3, 7.2, 4.6 Hz, 6H), 3.67 (d, J = 1.1 Hz, 3H), 2.98 (dd, J = 5.1, 2.7 Hz, 2H), 2.84 (td, J = 5.4,
5.0, 2.1 Hz, 5H), 2.42-2.35 (m, 5H), 1.98 (td, J = 2.8, 1.0 Hz, 1H).
13
C NMR (126 MHz,
Chloroform-d) δ 173.52, 130.07, 129.27, 128.61, 128.16, 128.08, 127.90, 127.39, 124.03, 82.55,
68.13, 51.56, 34.00, 31.92, 29.70, 25.63, 22.80, 16.88.

(4Z,7Z,10Z,13Z)-methyl 23-(trimethylsilyl)tricosa-4,7,10,13-tetraen-16,19,22-triynoate (2.8).
Compound 2.8 was synthesized similarly to compound 2.3, and collected as light yellow oil in
COOMe
COOMe
TMS
32
26% yield (17 mg).
1
H NMR (600 MHz, Chloroform-d) δ
H
5.46-5.34 (m, 8H), 3.68 (s, 3H), 3.20
(t, J = 2.4 Hz, 2H), 3.14 (p, J = 2.4 Hz, 2H), 2.95 (dt, J = 5.2, 2.4 Hz, 2H), 2.86-2.81 (m, 6H),
2.43-2.35 (m, 4H), 0.16 (s, 9H).
13
C NMR (151 MHz, CDCl
3
) δ 173.52, 129.54, 129.30, 128.52,
128.16, 128.13, 127.92, 127.57, 124.69, 99.86, 85.09, 78.75, 75.12, 73.83, 73.75, 51.55, 36.11,
34.03, 29.08, 25.65, 22.83, 18.75, 17.18, 11.42, -0.11.

(4Z,7Z,10Z,13Z,16Z,19Z)-methyl tricosa-4,7,10,13,16,19-hexaen-22-ynoate (2.9). Compound
2.9 was synthesized similarly to compound 2.4, and collected as light yellow oil in 18% yield
over two steps (43 mg).
1
H NMR (600 MHz, Methanol-d
4
) δ
H
5.46-5.35 (m, 12H), 3.65 (s, 3H),
2.96 (ddt, J = 5.0, 2.7, 0.6 Hz, 2H), 2.90-2.84 (m, 10H), 2.39-2.37 (m, 4H), 2.23 (t, J = 2.8 Hz,
1H).
13
C NMR (151 MHz, Methanol-d
4
) δ 175.26, 130.81, 130.32, 129.58, 129.21, 129.18,
129.15, 129.11, 129.07, 128.99, 128.50, 125.49, 83.18, 69.29, 52.06, 34.85, 26.60, 26.59, 26.58,
26.51, 26.43, 23.84, 17.46.

sodium (4Z,7Z,10Z,13Z,16Z,19Z)-tricosa-4,7,10,13,16,19-hexaen-22-ynoate (2.1). To 0.25
mL of a 2.28 M NaOH solution in ATP-ase free water, was added 0.25 mL of ethanol. The
carboxylic ester (2.28 mg, 6.47 mmol) was added to this solution and mixed well, without a
COOMe
COO
-
Na
+
33
stirbar. The reaction was kept at 0
o
C for 48 hours. The final product was purified by HPLC run
at 20% water:methanol. The title compound (2.1, 0.36 mg, 1.00 mmol) was collected as a white
solid in 15% yield.
1
H NMR (600 MHz, Methanol-d
4
) δ
H
5.50-5.27 (m, 12H), 2.96 (ddt, J = 5.0,
2.6, 0.6 Hz, 2H), 2.90-2.83 (m, 10H), 2.40-2.34 (m, 2H), 2.23 (t, J = 2.8 Hz, 1H), 2.21-2.17 (m,
2H).

13
C NMR (151 MHz, Methanol-d
4
) δ 178.89, 130.84, 130.83, 129.60, 129.57, 129.29,
129.26, 129.10, 129.05, 128.94, 128.90, 128.49, 125.47, 111.44, 69.27, 49.85, 39.13, 26.56,
26.42, 25.63, 17.46. LC-MS: calculated 337.21 [M-Na
+
]
-
, observed 337.2 [M-Na
+
]
-
.

(4Z,7Z,10Z,13Z,15E,19Z)-17-hydroxydocosa-4,7,10,13,15,19-hexaenoate (2.10). To 10 mL of
0.05 M sodium borate buffer at 0
o
C in a round bottom flask with a stirbar was added 0.328 mg
(0.001 mmol) of docosahexaenoic acid. Next, 13.1 Ku of 15-lipoxygenase was added and the
reaction was stirred at 0
o
C for 30 minutes in the open air. Sodium borohydride (approximately
20 mg) was added to the mixture, to reduce the peroxide to a hydroxyl group. Gas evolved and
the reaction foamed. The reaction was stirred for another 20 minutes at 0
o
C, and then the pH
was adjusted to approximately 4 by adding glacial acetic acid dropwise. The aqueous layer was
extracted with diethyl ether (3 x 10 mL), combined, dried over magnesium sulfate, filtered, and
then concentrated in vacuo. The crude product was analyzed by LC-MS/MS without further
purification. LC-MS/MS: calculated 343.23 [M-1], observed 343.2 [M-1].

COO
-
OH
34

(4Z,7Z,10Z,13Z,15E,19Z)-17-hydroxytricosa-4,7,10,13,15,19-hexaen-22-ynoate (2.11). To
2 mL of 0.05 M sodium borate buffer at 0
o
C in a round bottom flask with a stirbar was added
0.07 mg (0.0002 mmol) of 2.1. Next, 2.6 Ku of 15-lipoxygenase was added and the reaction was
stirred at 0
o
C for 30 minutes in the open air. Sodium borohydride (approximately 10 mg) was
added to the mixture, to reduce the peroxide to a hydroxyl group. Gas evolved and the reaction
foamed. The reaction was stirred for another 20 minutes at 0
o
C, and then the pH was adjusted to
approximately 4 by adding glacial acetic acid dropwise. The aqueous layer was extracted with
diethyl ether (3 x 10 mL), combined, dried over magnesium sulfate, filtered, and then
concentrated in vacuo. The crude product was analyzed by LC-MS/MS without further
purification. LC-MS/MS: calculated 353.21 [M-1], observed 353.3 [M-1].

COO
-
OH
35
2.5 Chapter 2 References

1
Singh, M. Indian J. Pediatr. 2005, 72, 239.

2
Petasis, N. A.; Akritopoulou-Zanze, I.; Fokin, V. V.; Bernasconi, G.; Keledjian, R.; Yang, R.;
Uddin, J.; Nagulapalli, K. C.; Serhan, C. N. Prostaglandins Leukot. Essent. Fatty Acids 2005, 73,
301.

3
Arita, M.; Bianchini, F.; Aliberti, J.; Sher, A.; Chiang, N.; Hong, S.; Yang, R.; Petasis, N. A.;
Serhan, C. N. J. Exp. Med. 2005, 201, 713.

4
Spite, M.; Norling, L. V.; Summers, L.; Yang, R.; Cooper, D.; Petasis, N. A.; Flower, R. J.;
Perretti, M.; Serhan, C. N. Nature 2009, 461, 1287.

5
Serhan, C. N.; Gotlinger, K.; Hong, S.; Lu, Y.; Siegelman, J.; Baer, T.; Yang, R.; Colgan, S. P.;
Petasis, N. A. J. Immunol. 2006, 176, 1848.

6
Serhan, C. N.; Dalli, J.; Karamnov, S.; Choi, A.; Park, C.; Xu, Z.; Ji, R.; Zhu, M.; Petasis, N.
A. FASEB J. 2012, 26, 1755.

7
Marcheselli, V. L.; Hong, S.; Lukiw, W. J.; Tian, X. H.; Gronert, K.; Musto, A.; Hardy, M.;
Gimenez, J. M.; Chiang, M.; Serhan, C. N.; Bazan, N. G. J. Biol. Chem. 2003, 278, 43807.

8
Mukherjee, P. K.; Marcheselli, V. L.; Serhan, C. N.; Bazan, N. G. Proc. Natl. Acad. Sci.
U. S. A. 2004, 101, 8491.

9
Serhan, C. N.; Petasis, N. A. Chem. Rev. 2011, 111, 5922.

10
Finaldi, A.-M.; Petasis, N. A. 243
rd
ACS National Meeting and Exposition, San Diego, CA,
March 2012.

11
Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B. Angew. Chem. Int. Ed. 2002,
41, 2596.

12
Wang, Q.; Chan. T. R.; Hilgraf, R.; Fokin, V. V.; Sharpless, K. B.; Finn, M. G. J. Am. Chem.
Soc. 2003, 125, 3192.

13
Yap, M. C.; Kostiuk, M. A.; Martin, D. D. O.; Perinpanayagam, M. A.; Hak, P. G.; Siddam,
A.; Majjigapu, J. R.; Rajaiah, G.; Keller, B. O.; Prescher, J. A.; Wu, P.; Bertozzi, C. R.; Falck, J.
R.; Berthiaume, L. G. J. Lipid Res. 2010, 51, 1566.

14
Baskin, J. M.; Prescher, J. A.; Laughlin, S. T.; Agard, N. J.; Chang, P. V.; Miller, I. A.; Lo, A.;
Codelli, J. A.; Bertozzi, C. R. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 16793.

36

15
Chang, P. V.; Prescher, J. A.; Sletten, E. M.; Baskin, J. M.; Miller, I. A.; Agard, N. J.; Lo, A.;
Bertozzi, C. R. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 1821.

16
Sandri, J.; Viala, J. J. Org. Chem. 1995, 60, 6627.

17
Dangi, B.; Obeng, M.; Nauroth, J. M.; Teymourlouei, M.; Needham, M.; Raman, K.;
Arterburn, L. M. J. Biol. Chem. 2009, 284, 14744.
37
Chapter 3. Design and Synthesis of Lipoxin Analog Building Blocks
3.1 Introduction
Lipoxins (LX) were first discovered and isolated in 1984.
1,

2
Lipoxins are derived from
arachidonic acid (AA), a polyunsaturated omega-6 fatty acid found in abundance in cell
membranes. Lipoxins contain a very specific trihydroxytetraene moiety that plays a critical role
in their ability to achieve signaling and resolution of inflammation.
2
There are two native
lipoxins, Lipoxin A
4
(LXA
4
) and Lipoxin B
4
(LXB
4
), both of which are formed through
oxygenation of AA by lipoxygenase enzymes.
3
Specifically, three different lipoxygenases (5-
lipoxygenase (5-LO), 12-lipoxygenase (12-LO), and 15-lipoxygenase (15-LO)) work together
through several pathways to yield both LXA
4
and LXB
4
.
3
Aspirin-acetylated cyclooxygenase-2
(COX-2) can produce the 15-epi versions of each of these molecules, so named the “aspirin-
triggered lipoxins” (ATL), shown in Figure 3.1.
3, 4

Figure 3.1- Structures of potent anti-inflammatory agents, lipoxins (LXA
4
and LXB
4
) and
aspirin-triggered lipoxins (15-epi-LXA
4
and 15-epi-LXB
4
).
3, 4

COOH
OH HO
OH
Lipoxin A
4
(LXA
4
)
OH HO
OH
COOH
Lipoxin B
4
(LXB
4
)
COOH
OH HO
OH
15-epi-LXA
4
OH HO
OH
COOH
15-epi-LXB
4
38
Although lipoxins are potent anti-inflammatory agents, they are short-lived due to their
ease of oxidation, reduction, and degradation in vivo. Isomerization of the C
11
-C
12
cis double
bond to the trans isomer, as well as epimerization at C
15
from R to S significantly reduces the
activity of the molecule.
5,

6,

7
These metabolomic processes deactivate the molecule and nullify
any of its valuable properties. Much effort and intensive studies have been performed to develop
metabolically stable analogs of lipoxins. Many previous studies involved the total synthesis of
lipoxin analogs to exhibit improved resistance to oxidation and longer metabolic half-lives in
vivo.
4, 8, 9, 10, 11, 12
Figure 3.2 shows a select number of active lipoxin analogs that were designed
specifically for improved stability compared with LXA
4
and 15-epi-LXA
4
. ZK-142 and ZK-994
(Figure 3.2) replaced C
3
with an oxygen atom, to prevent subsequent β-oxidation at carbons 2
and 3.
4, 9, 11, 12
The para-fluorophenoxy moiety of 16-parafluorophenoxy-15-epi-LXA
4
showed
enhanced stability to the C
15
end of the molecule as well as retaining bioavailability and activity
in vivo.
13, 14, 15

Figure 3.2- Select lipoxin analogs that have exhibited high anti-inflammatory potency and
enhanced metabolic stability.
COOMe
OH HO
OH
o-[9,12]-Benzo-15-epi-LXA
4
methyl ester
ZK-142
COOMe
OH HO
OH
16-parafluorophenoxy-15-epi-LXA
4
O F
O COOH
OH HO
OH
O F
ZK-994
O COOH
OH HO
OH
O F
39
In 2008, Petasis et. al. developed benzo-lipoxin derivatives, designed for improved
stability to the E, Z, E-triene moiety by replacing it with a benzene ring.
10, 16
Not only did this
increase the metablic stability of the molecule, and alleviated the possibility of isomerization at
the cis double bond, but it also allowed for a simplified total synthesis of lipoxin analogs.
Figure 3.2 shows o-[9,12]-benzo-15-epi-LXA
4
methyl ester which was synthesized by Petasis et.
al. and was found to have comparable anti-inflammatory action to LXA
4
, while possessing
enhanced metabolic stability.
10

Lipoxin analogs (ZK-994, as well as others) have already
undergone clinical trials for inflammatory disorders.
9,

17
Even though LXA
4
and ATL
derivatives show promise for as treatments for inflammatory disorders, more investigations are
needed to establish structure-activity relationships.

3.2 Results and Discussion
3.2.1 Design of Lipoxin Analog Building Blocks
We sought to design building blocks that could be used for the synthesis of novel lipoxin
and benzo-lipoxin analogs. Three functional groups were of particular interest to us: i.) para-
fluorphenoxy ii.) 3-oxa lipoxin A
4
diol (C
1
-C
8
) and iii.) benzo-lipoxins. These three groups of
building blocks were chosen because their functionality has already shown enhanced metabolic
stability in vivo. In order for the building blocks to be useful, we needed them to have some
“handle” or functionality that would make them easy to couple to other pieces. For this reason,
we chose to synthesize aldehydes, alkynes, and vinyl halides due to their versatility and broad
application to coupling reactions. We also needed to synthesize the pieces in high enantiomeric
excess at relevant stereocenters. The hydroxyl stereocenters of potent lipoxin derivatives are of
40
utmost importance. Any presence of the inactive epimer would decrease the anti-inflammatory
activity as well as skew the biological testing data.
The first group of building blocks we designed was para-fluorophenoxy pieces 3.1 and
3.2, shown in Figure 3.3. We designed a strategy to use chiral starting materials and thereby
avoid challenging stereoselective reactions, which often result in production of some quantity of
undesired isomer. The para-fluorophenoxy pieces can be synthesized starting from
commercially available S-glycidol, shown in Figure 3.3. Vinyl iodide 3.1 is a valuable molecule
because it can easily be coupled to an aromatic ring via palladium-mediated Suzuki or Heck
coupling. Alkyne 3.2 could be used in coupling reactions, or alternatively the alkyne could be
transformed into another useful functional group.

Figure 3.3- Retrosynthesis of para-fluorophenoxy building blocks 3.1 and 3.2 from a chiral
starting material.

The naturally occurring sugar, L-rhamnose, served as a chiral pool for the 3-oxa-diol
pieces, shown in Figure 3.4. The vinyl iodides 3.3 and 3.4 could be synthesized by utilizing the
pre-existing stereochemistry of L-rhamnose and opening the ring. Different protecting groups
I
OTBDPS
O F
3.1
OTBDPS
O F
3.2
O
OH
S-Glycidol
41
were installed to protect the diol and the carboxylic acid, allowing for possible selective
deprotection of one over the other if necessary. We also alternated between the bulkiness of the
protecting groups the protecting groups on 3.3 and 3.4. Notably, 3.3 has a bulky t-butyl ester
group, and a smaller dimethyl acetal protecting group on the diol, while 3.4 has a small methyl
ester and a bulky cyclohexane acetal group on the diol. Differentiation between the protecting
groups is a key element to pieces 3.3 and 3.4, so that later we may selectively deprotect one
group over the other, if necessary.

Figure 3.4- Retrosynthesis of 3-oxa lipoxin A
4
building blocks 3.3 and 3.4, derived from the
chiral sugar L-Rhamnose.

The last group of builiding blocks we designed was benzo-lipoxin building blocks. For
these pieces we Suzuki coupled vinyl iodide 3.1 to different commercially available
bromophenylboronic acids. By varying the substitution of the bromine on the phenylboronic
acid, we were able to control the positioning of the substitution on the benzene ring.
O OH
HO
OH
OH
L-Rhamnose
O
O O O
O
I
3.3
O
O O O
O
I
3.4
42
3.2.2 Synthesis of Fluorophenoxy Building Blocks

Scheme 3.1- The first part of the synthesis of para-fluorophenoxy building blocks 3.1 and 3.2.
Scheme 3.1 and Scheme 3.2 detail the synthesis of two building blocks vinyl iodide (3.1)
and alkyne (3.2). First, S-glycidol underwent a ring-opening reaction catalyzed by cesium
fluoride to produce diol 3.5 in decent yield. Next, 3.5 was protected under mild conditions using
TBS-Cl to selectively silyl-protect the primary alcohol. Singly protected alcohol 3.6 could then
be protected at the secondary alcohol with TBDPS-Cl, using the same conditions as the previous
protection, again in good yield. With the fully protected 3.7 in hand, a careful selective
deprotection could be performed using camphor sulfonic acid to remove the TBS functional
group on the primary alcohol. The selective deprotection was monitored via TLC very carefully
to avoid deprotecting both of the alcohols, once the reaction was completed it was quenched with
triethyl amine. Compound 3.8 was formed in decent yield and in high purity.
HO F
O
(S) (S)
OH
a
HO
OH
O F
+
TBSO
OH
O F
b
c
d
TBSO
OTBDPS
O F
HO
OTBDPS
O F
3.5
3.6
3.7
3.8
Reagents and condtions: (a) CsF (cat.), DMF, 90
o
C, 2 days, 66% (b) TBSCl, DMAP, Imidazole, DCM, rt, overnight, 82%
(c) TBDPSCl, DMAP, Imidazole, DCM, rt, overnight, 84% (d) Camphor sulfonic acid, DCM:MeOH (1:1), rt, 30 min. then
TEA, 69%
43

Scheme 3.2- The second half of the synthesis of vinyl iodide (3.1) and alkyne (3.2) para-
fluorophenoxy building blocks.
A radical oxidation reaction converted alcohol 3.8 to the aldehyde 3.9 at 0
o
C in only 20
minutes.
18
The protected para-fluorophenoxy aldehyde 3.9 is a very useful intermediate for
adding diverse functionality to the para-fluorophenoxy building blocks because the aldehyde can
easily be converted into other functional groups. From aldehyde 3.9, vinyl iodide 3.1 can be
synthesized directly via a Takai reaction in moderate yield.
19
The fluorophenoxy aldehyde 3.9
can also undergo a Ramirez-Corey-Fuchs reaction to ultimately produce a terminal alkyne.
20,

21

First, 3.9 was reacted with CBr
4
and PPh
3
at low temperature to produce the 1,1’-dibromoolefin
in 73% yield. Second, n-butyl lithium was added to 3.10 at -78
o
C to result in the terminal
alkyne (3.2) formation in good yield. The para-fluorophenoxy vinyl iodide (3.1) and para-
fluorophenoxy alkyne (3.2) were both successfully synthesized.

OTBDPS
O F
O
OTBDPS
O F
I
OTBDPS
O F
Reagents and condtions: (a) TEMPO, Trichloroisocyanuric acid, DCM, 0
o
C 20 min. then rt 20 min., 50% (b) CrCl
2
,
Iodoform, THF, 0
o
C 3hrs, then rt for 1 hr, 69% (c) CBr
4
, PPh
3
, DCM, 0
o
C 40 min. then -78
o
C, then 0
o
C for 1 hr, 73%
(h) n-BuLi (2.5 M in Hexanes), Diethyl Ether, -78
o
C for 20 min. then 0
o
C for 20 min., 74%
HO
OTBDPS
O F
a
b
Br
OTBDPS
O F
Br
c
d
3.8
3.9
3.1
3.10
3.2
44
3.2.3 Synthesis of 3-Oxa Lipoxin A
4
Analog Building Blocks
3-Oxa lipoxin A
4
analog building blocks similar in structure to 3.3 and 3.4 have
previously been reported.
9
However, the pieces synthesized herein are novel due to their vinyl
iodide functionality. Scheme 3.3 illustrates the synthesis of vinyl iodide 3.3 in five total steps.
Beginning with L-rhamnose as a chiral starting material, the cis-diol at C
3
and C
4
were protected
by forming a five membered ring and producing dimethyl acetal 3.11.
22
The five membered ring
acetal can only be formed with the cis-diol, which is an important feature of this synthesis
because it allows for differentiation between the four hydroxyl groups of the sugar. A reductive
ring-opening using sodium borohydride allowed for a smooth transformation from 3.11 to 3.12.
Next, the primary alcohol of the triol (3.12) was selectively reacted in an SN
2
reaction with t-
butyl bromoacetate yielding ether 3.13, in low yield.
9
The low yield may be attributed to the
mild condtions of the reaction as well as the poor selectivity. However, selectivity of the SN
2

reaction to product 3.13 is not a major issue because the undesired side products will not be
reactive in the next step of the synthesis.
45

Scheme 3.3- The synthesis of 3-oxa vinyl iodide building block 3.3.
Taking advantage of the vicinal diol functionality on 3.13, aldehyde 3.14 could be
produced using sodium periodate and silica in dichloromethane and water.
23
Periodate requires
the vicinal diol in order to form a five-membered ring and oxidize the alcohol to the aldehyde.
Aldehyde 3.14 was produced in low yield, but was easily separable from the starting material
(3.13) due to the high polarity of the diol functional group. From aldehyde 3.14, a Takai reaction
was proformed to construct the vinyl iodide (3.3) in 37% yield.

O OH
HO
O
O
O OH
HO
OH
OH
a b
HO OH
OH
O O
c
HO OH
O
O O O
O
d
O
O
O O O
O
e
O
O O O
O
I
Reagents and conditions: (a) 2-Methoxypropene, p-Toluenesulfonic acid, Drierite, DMF, 0
o
C for 3 hrs, then rt overnight,
93% (b) NaBH
4
, MeOH, 0
o
C for 3 hrs, then rt overnight, 64% (c) t-Butyl bromoacetate, NaOH, TBAS, Toluene, rt,
overnight, 25% (d) NaIO
4
, SiO
2
, DCM, H
2
O, rt 2 hrs., 36% (e) CrCl
2
, Iodoform, THF, 0
o
C 3hrs, then rt for 1 hr, 37%
L-Rhamnose
3.11
3.12
3.13
3.14
3.3
46

Scheme 3.4- The synthesis of 3-oxa vinyl iodide 3.4, similar to the synthesis of 3.3, but with
different protecting groups.
A similar synthetic route as shown in Scheme 3.3 was followed for the synthesis of vinyl
iodide 3.4 (shown in Scheme 3.4). Again, the synthesis begins with the protection of L-
rhamnose, however the protecting group is now a cyclohexane acetal (3.15). The protection
reaction gave the product (3.15) in good yield. A ring opening reaction with sodium borohydride
produced the triol (3.16), which was used crude without purifcation. Next, an SN
2
reaction was
attempted with methyl bromoacetate and 3.16 to synthesize the ether 3.17 similarly to how 3.13
O OH
HO
O
O
O OH
HO
OH
OH
a b
HO OH
OH
O O
c
HO OH
O
O O O
O
d
O
O
O O O
O
e
O
O O O
O
I
Reagents and conditions: (a) 1,1-dimethoxycyclohexane, p-Toluenesulfonic acid, Drierite, DMF, 0
o
C for 3 hrs, then rt
overnight, 70% (b) NaBH
4
, MeOH, 0
o
C for 3 hrs, then rt overnight (c) Methyl bromoacetate, t-BuOK, THF, 0
o
C for 3 hrs,
then rt overnight, 63% (d) NaIO
4
, SiO
2
, DCM, H2O, rt 2 hrs., 17% (e) CrCl
2
, Iodoform, THF, 0
o
C 3hrs, then rt for 1 hr,
49%
L-Rhamnose
3.15
3.16
3.17
3.18
3.4
47
was attained. This reaction was found to be problematic, even more so than the one shown in
Scheme 3.3 where t-butyl bromoacetate was used for the same reaction. We believe that the
bulky t-butyl group enhanced the selectivity of this reaction for the primary alcohol over the
secondary alcohols, and thus when switching to the methyl bromoactate, the reaction resulted in
an extremely low yield and a mixture of all three possible products. When we altered the
conditions to use a less nucleophilic base, t-BuOK and the appropriate solvent (THF), a yield of
63% was observed, but this was still a mixture of the three possible ether products, which were
inseprable.
24
However, the undesired side-products would be unreactive in the next step of the
synthesis, which requires a 1,2-diol to form the aldehyde (3.18). The synthesis was continued
with 3.17 containing the mixture of inseprable side-products.
Aldehyde (3.18) was attained using the same method as for producing 3.14, with sodium
periodate bonding to the vicinal diol, but in only 17% yield. The very low yield can be attributed
to the presence of impurities and side-products in with the starting material, which would cause
the weight of the starting material to be lower than measured and thus the yield would be lower
than expected. A Takai reaction of 3.18 with chromium(II) chloride and iodoform produced the
vinyl iodide product (3.4) in 49% yield. We successfully achieved the five-step synthesis of 3.4,
as a building block for lipoxin analogs.
We wanted to perform a Wittig reaction to extend the chain of aldehyde 3.14 to
synthesize some lipoxin analogs similar to ZK-994 (see Scheme 3.5). Fortunately, a past
research group member (Jasim Uddin) previously synthesized a large quantity of phosphonium
salt (3.19, shown in Scheme 3.5), and allowed us to use some of his pure compound for this
purpose. We successfully performed the Wittig reaction coupling the aldehyde 3.20 to the
phosphonium salt (3.19) using n-butyl lithium at low temperature, and in 62% yield. We were
48
very pleased to produce 3.20 in decent yield, using this challenging Wittig reaction. The product
obtained (3.20) was a mixture of cis and trans isomers that were separable via silica column
chromatography. However, we also attempted to isomerize the undesired cis isomer side-
product to the trans isomer (3.21) using a radical reaction of iodine and light. Although this
isomerization method appeared promising, the reaction yielded no product, only decomposition.
Due to the failure of the isomerization reaction, we decided to use column chromatography to
isolate 3.21 rather than jeopordize decomposing any more of the precious mixture (3.20).

Scheme 3.5- A Wittig reaction to couple the aldehyde (3.14) with the phosphonium salt (3.19) to
produce 3.20, and the attempted isomerization of the cis/trans mixture (3.20) to 3.21.

TMS
PPh
3
Br
O
O
O O O
O
+
a
O
O O O
O
TMS
b
O
O O O
O
TMS
Reagents and conditions: (a) n-BuLi (2.5 M in Hexanes), THF, -78
o
C, 0
o
C, -78
o
C, then rt for 3hrs., 62% (b) I
2
, DCM, hv,
rt overnight, reaction failed
3.19
3.14
3.20
3.21
49
3.2.4 Synthesis of Benzo-Lipoxin Building Blocks
We envisaged assembling novel benzo-lipoxins starting from the fluorophenoxy bottom
piece previously synthesized (see Scheme 3.6). Knowing that the Suzuki reaction can be a
powerful tool in the assembly of benzo-lipoxins, we designed the construction of our novel
benzo-lipoxin analogs that employed Suzuki coupling reactions. Herein, we utilized the para-
fluorophenoxy vinyl iodide (3.1) and Suzuki coupled it to the commercially available 3-
bromophenylboronic acid directly.
25, 26
The catalyst used for the Suzuki coupling was
tetrakis(triphenylphosphine)palladium(0) and the reaction was heated overnight at 90
o
C. The
stereoselective-coupling product (3.22) was produced in 25% yield.

Scheme 3.6- Suzuki coupling and Miyaura borylation reactions to produce the meta-benzo-
lipoxin building blocks.
The aryl bromide (3.22) was then subjected to a Miyaura borylation reaction to convert
the bromide into a boronic ester (3.23).
27
This reaction needed heat to proceed, and beared the
I
OTBDPS
O F
B(OH)
2
Br
+
a
OTBDPS
O F
Br
b
OTBDPS
O F
BPin
3.1
3.22
3.23
Reagents and conditions: (a) Pd(PPh
3
)
4
, K
2
CO
3
, Dioxane, H
2
O, 90
o
C, overnight, 25% (b) PdCl
2
(dppf),
Bis(pinacolato)diboron, DMSO, Dioxane, 80
o
C, 2 hrs., 21%
50
product in low yield. Overall, in two steps we successfully made a functionalized meta-benzo-
lipoxin building block that can easily undergo another Suzuki coupling reaction to attach any
number of top chain pieces, particularly a vinyl iodide such as 3.3 or 3.4.
The same synthetic route can be applied to the para-benzo-lipoxin building block, shown
in Scheme 3.7. Beginning with commercially available 4-bromophenylboronic acid, a Suzuki
coupling reaction was performed with vinyl iodide 3.1 in 33% yield. It is probable that the slight
increased in yield of this Suzuki coupling reaction, compared with that of Scheme 3.6, is due to
the difference in benzene ring substitution. The para-phenylboronic acid has less steric bulk
compared to that of the meta-substituted phenylboronic acid and thus could result in a slightly
higher yield of product. With 3.24 in hand, we performed a Miyaura borylation to aquire the
boronic ester building block (3.25).

Scheme 3.7- Suzuki coupling and Miyaura borylation reactions to produce the para-benzo-
lipoxin building blocks.
I
OTBDPS
O F
Br
+
a
OTBDPS
O F
b
OTBDPS
O F
B(OH)
2
Br
PinB
Reagents and conditions: (a) Pd(PPh
3
)
4
, K
2
CO
3
, Dioxane, H
2
O, 90
o
C, overnight, 33% (b) PdCl
2
(dppf),
Bis(pinacolato)diboron, DMSO, Dioxane, 80
o
C, 2 hrs., 19%
3.1
3.24
3.25
51
3.3 Conclusion
Several lipoxin analog building blocks were synthesized successfully for later assembly
to final compounds. A para-fluorophenoxy vinyl iodide building block (3.1) was synthesized in
six steps, and high purity. An acetylenic para-fluorophenoxy piece (3.2) was also completed
successfully in seven overall steps. Two 3-oxa-vinyl iodide building blocks with varying
protecting groups (3.3 and 3.4) were designed and made. An aldehyde intermediate (3.14) was
used to assemble 3.20, a major component of biologically active ZK-994 and similar analogs. A
Suzuki coupling and then a Miyaura borylation reaction were used to synthesize the aryl boronic
esters 3.23 and 3.25. We now have several pieces prepared and further studies are underway to
utilize the pieces synthesized herein.

3.4 Experimental
All reactions, unless otherwise noted, were carried out with commercially available
starting materials and solvents, which were used without further purification. The majority of
the chemicals were purchased from Sigma Aldrich, GFS chemicals, TCI America, and VWR
International. All glassware and reaction vessels used were standard and commercially
available. Reactions performed under “inert atmosphere” implies the reaction was run in a round
bottom flask affixed with a three-way adapter and an argon balloon, or the flask was affixed with
a septum which was punctured with a needle affixed to an argon balloon.
1
H and
13
C NMR
spectra were collected using a Varian Mercury 400, Varian 400-MR, Varian 500, or Varian 600
NMR spectrometers, using residual
1
H and
13
C peaks of deuterated solvents as internal standards.
Manual flash column chromatography was run with silica gel purchased from Sorbent
52
Technologies (60 Å, 40-63 µm). A Biotage Isolera One was used for automated flash column
chromatography, with standard Biotage silica cartridges (60 Å) for purification. LC-MS analysis
was performed on an Agilent Technologies 1200 infinity series instrument.

Procedures

2S, 3-(4-fluorophenoxy)propane-1,2-diol (3.5). To a flame-dried flask under inert atmosphere,
equipped with a stir bar and three-way adapter, was added fluorophenol (4.09 g, 36.48 mmol)
and cesium fluoride (0.31 g, 2.04 mmol). Anhydrous DMF (15 mL) was added via syringe, and
then the mixture was stirred under argon at room temperature for 1 hour. S-Glycidol (2.7 mL,
40.67 mmol, 98% ee) was added via syringe, and then the reaction was stirred at 90
o
C for 2 days.
The reaction was cooled to room temperature, then quenched with distilled water (30 mL) and
extracted with ethyl acetate (3 x 100 mL). The combined organic layers were washed with brine,
dried with anhydrous magnesium sulfate, filtered and then concentrated in vacuo. The crude
product was purified by column chromatography using silica gel and a mobile phase of ethyl
acetate: hexanes (1: 1). Compound 3.5 was isolated in decent yield, 5.01 g (66%).
1
H NMR
(400 MHz, Chloroform-d) δ
H
6.93 (m, 2H), 6.86 (dd, J = 9.1, 4.1 Hz, 2H), 4.11 (quintet, J = 5.8
Hz, 1H), 3.96 (m, 2H), 3.85 (dd, J = 11.4, 3.9 Hz, 1H), 3.75 (dd, J = 11.4, 5.4 Hz, 1H).

53

2R-1-(t-butyldimethylsilyloxy)-3-(4-fluorophenoxy)propan-2-ol (3.6). Under inert
atmosphere, TBSCl (4.06 g, 26.94 mmol), DMAP (0.17 g, 1.35 mmol), and imidazole (1.84 g,
27.03 mmol) were added to a flame-dried flask with a stir bar and 3-way adapter. Anhydrous
DCM (60 mL) was added via syringe, and the mixture was cooled to 0
o
C. A solution of the diol
(3.5, 5.01 g, 26.91 mmol) in 20 mL anhydrous DCM was cannulated into the reaction flask, after
which the reaction was warmed to room temperature and stirred under argon overnight. The
reaction was quenched with ammonium chloride, extracted with diethyl ether (3 x 100 mL),
washed with brine, dried with anhydrous magnesium sulfate, filtered, and then concentrated in
vacuo. The product was purified with silica gel by flash chromatography using a 10% ethyl
acetate: hexanes mobile phase. The purified product (6.68 g, 82%) was obtained in good yield.
1
H NMR (400 MHz, Chloroform-d) δ
H
6.97 (dd, J = 9.2, 8.2 Hz, 2H), 6.85 (dd, J = 9.2, 4.3 Hz,
2H), 4.01 (quintet, J = 5.6 Hz, 1H), 3.96 (d, J = 5.4 Hz, 2H), 3.71 (m, 2H), 0.90 (s, 9H), 0.08 (s,
3H), 0.07 (s, 3H).

2R-1-(t-butyldimethylsilyloxy)-2-(t-butyldiphenylsilyloxy)-3-(4-fluorophenoxy)
propane (3.7). Under inert atmosphere, TBDPSCl (6.95 mL, 26.73 mmol), DMAP (1.36 g,
11.13 mmol), and imidazole (1.82 g, 26.73 mmol) were stirred in anhydrous DCM (90 mL) at
0
o
C. A solution of the alcohol (3.6, 6.68 g, 22.23 mmol) in dry DCM (10 mL) was cannulated
54
into the reaction mixture, after which the reaction was warmed to room temperature and stirred
overnight. The reaction was quenched with ammonium chloride, extracted with diethyl ether (3
x 100 mL), washed with brine, dried with anhydrous magnesium sulfate, filtered, and then
concentrated in vacuo. The product was purified with silica gel by flash chromatography using a
2% ethyl acetate: hexanes mobile phase, and 9.77 g (84%) of purified product was obtained.
1
H
NMR (400 MHz, Chloroform-d) δ
H
7.66 (m, 4H), 7.46-7.28 (m, 6H), 6.89 (dd, J = 9.1, 8.3 Hz,
2H), 6.67 (dd, J = 9.1, 4.3 Hz, 2H), 4.08-3.96 (m, 1H), 3.87 (dd, J = 9.6, 5.7 Hz, 1H), 3.67-3.56
(m, 2H), 1.06 (s, 9H), 0.82 (s, 9H), -0.05 (s, 3H), -0.09 (s, 3H).

2S-2-(t-butyldiphenylsilyloxy)-3-(4-fluorophenoxy)propan-1-ol (3.8). The doubly protected
starting material (3.7, 9.77 g, 18.69 mmol) was stirred at room temperature in a solution of (1:1)
DCM: MeOH (66 mL). Camphor sulfonic acid (4.34 g, 18.68 mmol) was added to the mixture,
and stirred for 30 minutes, before being monitored by TLC for reaction completion. The
reaction was then quenched by triethylamine (2.61 mL, 18.71 mmol) and then concentrated in
vacuo. Ammonium chloride was added; the solution was extracted with diethyl ether (3 x 100
mL), washed with brine, dried with anhydrous magnesium sulfate, filtered, and then concentrated
in vacuo. The product was purified with silica gel by flash chromatography using a 10% ethyl
acetate: hexanes mobile phase, and 5.48 g (69%) of purified product was obtained.
1
H NMR
(400 MHz, Chloroform-d) δ
H
7.75-7.65 (m, 4H), 7.50-7.32 (m, 6H), 6.87 (dd, J = 9.1, 8.2 Hz,
2H), 6.58 (dd, J = 9.1, 4.3 Hz, 2H), 4.11 (m, 1H), 3.94 (dd, J = 9.6, 6.3 Hz, 1H), 3.86 (dd, J =
9.5, 5.5 Hz, 1H), 3.70 (d, J = 3.8 Hz, 2H), 1.09 (s, 9H).

55

2R-2-(t-butyldiphenylsilyloxy)-3-(4-fluorophenoxy)propanal (3.9). A solution of the alcohol
(3.8, 0.50 g, 1.18 mmol) in anhydrous DCM (5 mL) was stirred at 0
o
C under argon atmosphere.
Trichloroisocyanuric acid (0.287 g, 1.23 mmol) was added to the solution, followed by TEMPO
(0.002 g, 0.0128 mmol) (CAUTION! Exothermic!). The reaction was warmed to room
temperature and allowed to stir for 20 minutes, monitoring by TLC until completion. The
reaction was filtered through celite, quenched with an aqueous solution of saturated sodium
bicarbonate, which was then extracted with diethyl ether (3 x 10 mL). The combined organic
layers were washed with brine, dried over anhydrous magnesium sulfate, filtered, and
concentrated in vacuo. Flash column chromatography purification (10% ethyl acetate: hexanes)
using silica gel afforded the title compound (0.252 g, 50%).
1
H NMR (400 MHz, Chloroform-d)
δ
H
9.78 (d, J = 0.7 Hz, 1H), 7.79-7.69 (m, 4H), 7.54-7.33 (m, 6H), 6.95 (dd, J = 9.1, 8.1 Hz, 2H),
6.73 (dd, J = 9.1, 4.3 Hz, 2H), 4.39 (ddd, J = 5.1, 4.0, 0.9 Hz, 1H), 4.18-4.05 (m, 2H), 1.19 (s,
9H).

2S, 3E, 1-(4-fluorophenoxy)-4-iodo-but-3-en-2-ol (3.1). To a solution of CrCl
2
(1.35 g, 10.94
mmol) in dry THF (10 mL) at 0
o
C, was cannulated iodoform (1.72 g, 4.37 mmol) in THF (10
mL). The aldehyde (3.9, 0.445 g, 1.05 mmol) in 6 mL of THF was added to the mixture via
cannula. The reaction was stirred at 0
o
C for 3 hours, followed by 1 hour at room temperature.
56
The reaction was quenched with distilled water and extracted with diethyl ether (3 x 25 mL).
The combined organic layers were washed with brine, dried over anhydrous magnesium sulfate,
filtered and concentrated in vacuo. Flash column chromatography purification (5% ethyl acetate:
hexanes) using silica gel afforded the desired product (0.4 g, 69%).
1
H NMR (500 MHz,
Chloroform-d) δ
H
7.73-7.61 (m, 4H), 7.48-7.30 (m, 6H), 6.93-6.84 (m, 1H), 6.62-6.55 (m, 2H),
6.22 (dd, J = 14.5, 1.6 Hz, 1H), 4.41 (qd, J = 5.9, 1.1 Hz, 1H), 3.81 (ddd, J = 37.3, 9.6, 5.9 Hz,
2H), 1.08 (d, J = 0.6 Hz, 9H).
13
C NMR (126 MHz, Chloroform-d) δ 158.18, 156.28, 154.41,
144.79, 135.83, 133.36, 132.98, 129.91, 127.68, 115.76, 115.58, 115.38, 79.02, 74.01, 71.38,
31.56, 26.93.

S-tert-butyl((4,4-dibromo-1-(4-fluorophenoxy)but-3-en-2-yl)oxy)diphenylsilane (3.10). To a
flame-dried flask under inert atmosphere, equipped with a stir bar and three-way adapter was
added CBr
4
(0.40 g, 1.21 mmol) and dry DCM (2 mL). The mixture was cooled to 0
o
C, and then
PPh
3
(0.63 g, 2.40 mmol) in DCM (2 mL) was added via cannula. The reaction was allowed to
stir for 30 minutes at 0
o
C, and then it was cooled to -78
o
C, and the aldehyde (3.9, 0.25 g, 0.59
mmol) in DCM (4 mL) was cannulated into the reaction mixture. The reaction was then
warmed to 0
o
C and stirred at that temperature for 1 hour. The reaction was quenched with
saturated aqueous sodium bicarbonate solution and extracted with DCM (3 x 10mL). The
combined organic layers were dried over anhydrous sodium sulfate, filtered, and concentrated in
vacuo. The crude product was purified on silica gel by flash chromatography (5% ethyl acetate:
hexanes), yielding 0.25 g (73%) of pure product.
1
H NMR (500 MHz, Chloroform-d) δ
H
7.68
57
(ddd, J = 16.0, 7.9, 1.4 Hz, 4H), 7.49-7.32 (m, 6H), 6.91 (dd, J = 9.1, 8.2 Hz, 2H), 6.70 (dd, J =
9.1, 4.3 Hz, 2H), 6.50 (d, J = 8.1 Hz, 1H), 4.65 (ddd, J = 8.0, 6.4, 4.6 Hz, 1H), 3.97-3.82 (m,
2H), 1.07 (s, 9H).
13
C NMR (126 MHz, Chloroform-d) δ 158.25, 156.36, 154.61, 137.87,
135.88, 135.86, 133.14, 133.06, 129.91, 129.84, 127.71, 127.64, 115.80, 115.62, 115.56, 115.49,
91.49, 72.87, 70.75, 26.87, 19.28.

S-tert-butyl((1-(4-fluorophenoxy)but-3-yn-2-yl)oxy)diphenylsilane (3.2). A solution of the
dibromoolefin (3.10, 0.25 g, 0.43 mmol) in anhydrous diethyl ether (4 mL) was cannulated into a
flame-dried reaction flask and cooled to -78
o
C. A 2.5 M solution of n-BuLi in hexanes (0.52
mL, 1.3 mmol) was added by syringe, and the reaction was stirred at -78
o
C for 20 minutes. The
temperature was then raised to 0
o
C, and the reaction was stirred for 20 minutes more. The
reaction was quenched with ammonium chloride, extracted with diethyl ether (3 x 10 mL),
washed with brine, dried with anhydrous magnesium sulfate, filtered, and then concentrated in
vacuo. Flash column chromatography purification (5% ethyl acetate: hexanes) using silica gel
afforded the desired product (0.135 g, 74%).
1
H NMR (500 MHz, Chloroform-d) δ
H
7.80-7.69
(m, 4H), 7.48-7.32 (m, 6H), 6.94-6.87 (m, 2H), 6.72-6.66 (m, 2H), 4.73-4.62 (m, 1H), 4.17-3.97
(m, 2H), 2.40 (dd, J = 2.1, 0.6 Hz, 1H), 1.09 (d, J = 0.6 Hz, 9H).
13
C NMR (126 MHz,
Chloroform-d) δ 158.31, 154.44, 136.07, 135.93, 135.55, 133.03, 132.96, 129.83, 127.59,
127.52, 115.81, 115.62, 114.58, 114.38, 81.94, 73.94, 72.43, 62.64, 26.83.

58

Methyl-2,3-O-isopropylidene-L-rhamnose (3.11). A solution of L-rhamnose (5.0 g, 27.45
mmol) and Drierite (1 g) in DMF (50 mL) were stirred at 0
o
C, under argon. 2-Methoxypropene
(5.25 mL, 54.82 mmol) was added via syringe followed by quick addition of p-toluenesulfonic
acid (0.020 g, 0.105 mmol). The reaction was stirred at 0
o
C for 3 hours, and then room
temperature overnight. The reaction was quenched with sodium carbonate (5 g), and allowed to
stir for one more hour. The reaction was then filtered through celite, rinsed with ethyl acetate,
and concentrated in vacuo to give a syrupy crude product. Flash column chromatography
purification (50% ethyl acetate: hexanes) using silica gel afforded the desired product (5.21 g,
93%) as a white solid.
1
H NMR (400 MHz, DMSO-d
6
) δ
H
6.29 (d, J = 4.3 Hz, 1H), 5.07 (d, J =
4.3 Hz, 1H), 4.70 (dd, J = 5.9, 3.4 Hz, 1H), 4.60 (d, J = 5.8 Hz, 1H), 4.41 (d, J = 5.9 Hz, 1H),
3.80 (dt, J = 8.6, 6.1 Hz, 1H), 3.65 (dd, J = 8.6, 3.4 Hz, 1H), 1.33 (s, 3H), 1.24 (s, 2H), 1.13 (d, J
= 6.2 Hz, 3H).

1-deoxy-4,5-O-(1-methylethylidene)-L-mannitol (3.12). Under inert atmosphere, anhydrous
NaBH
4
(3.08 g, 81.42 mmol) was added to a flame-dried flask, equipped with a stirbar and three-
way adapter. The flask was cooled to 0
o
C, and then anhydrous methanol (30 mL) was syringed
59
in. A solution of the diol (3.11, 5.21g, 25.51 mmol) in 10 mL anhydrous methanol was added to
the reaction flask via cannula. The reaction was stirred at 0
o
C for three hours, followed by
stirring at room temperature overnight. Acetic acid was added to the reaction until a pH of 6 was
reached. The reaction was then filtered through celite, rinsed with ethyl acetate and methanol,
and concentrated in vacuo. Flash column chromatography purification (pure ethyl acetate
followed by 30% methanol: ethyl acetate) using silica gel afforded the triol (3.27 g, 64%) as a
clear/slight yellow oil.
1
H NMR (400 MHz, Methanol-d
4
) δ
H
4.44 (dd, J = 6.9, 2.7 Hz, 1H), 4.23
(ddd, J = 6.9, 5.8, 4.8 Hz, 1H), 3.82-3.64 (m, 3H), 3.38 (dd, J = 8.0, 2.7 Hz, 1H), 1.47 (s, 3H),
1.36 (s, 3H), 1.25 (d, J = 6.2 Hz, 3H).
13
C NMR (126 MHz, Methanol-d
4
) δ 109.05, 79.22,
77.16, 74.43, 69.36, 62.29, 27.37, 25.29, 20.48.

Tert-butyl-2-(((4S,5R)-5-(1,2-dihydroxypropyl)-2,2-dimethyl-1,3-dioxolan-4-
yl)methoxy)acetate (3.13). The triol (3.12, 3.0 g, 14.55 mmol) was added to a round bottom
flask, with a stir bar and toluene (37 mL). Then tert-butyl bromoacetate (2.58 mL, 17.47 mmol)
was added to the solution, followed by an aqueous solution of NaOH (25% by wt., 3 mL). The
mixture then became cloudy white, and then tetra-butyl ammonium sulfate (0.35 g, 1.03 mmol)
was added in. The reaction flask was equipped with a septum and allowed to stir under argon at
room temperature overnight. The reaction was diluted with ethyl acetate, quenched with
saturated aqueous monobasic potassium phosphate, and extracted with ethyl acetate (3 x 100
60
mL). The combined organic layers were dried over anhydrous sodium sulfate, filtered, and
concentrated in vacuo. The crude product was purified by flash column chromatography over
silica, run at 50% ethyl acetate: hexanes. The purified product was obtained, although in low
yield (1.08 g, 25%).
1
H NMR (500 MHz, Chloroform-d) δ
H
4.47-4.39 (m, 1H), 4.37 (dd, J = 6.6,
3.5 Hz, 1H), 4.10-3.92 (m, 2H), 3.89-3.83 (m, 1H), 3.79 (p, J = 6.4 Hz, 1H), 3.68 (dd, J = 9.6,
5.3 Hz, 1H), 3.63 (dd, J = 6.1, 3.5 Hz, 1H), 1.48 (s, 3H), 1.48 (s, 9H), 1.37 (d, J = 0.8 Hz, 3H),
1.30 (d, J = 6.4 Hz, 3H).

Tert-butyl 2-(((4S,5S)-5-formyl-2,2-dimethyl-1,3-dioxolan-4-yl)methoxy)acetate (3.14). In a
round bottom flask, was added a stir bar, SiO
2
(4.64 g, 77.23 mmol) and DCM (39 mL). A
solution of NaIO
4
(1.41 g, 6.59 mmol) in H
2
O (8 mL) was added to the stirring mixture at room
temperature. The diol (3.13, 0.979 g, 3.35 mmol) in DCM (3 mL) was added to the reaction
flask, and the reaction was stirred at room temperature and monitored by TLC, until it was
completed after 2 hours of stirring. The reaction was filtered, dried with anhydrous sodium
sulfate, and then filtered again, before being concentrated in vacuo. Flash column
chromatography on the crude product (30% ethyl acetate: hexanes) yielded the purified aldehyde
(0.298 g, 36%).
1
H NMR (500 MHz, Chloroform-d) δ
H
9.71 (d, J = 2.2 Hz, 1H), 4.60 (dt, J =
8.2, 4.3 Hz, 1H), 4.45 (dd, J = 7.7, 2.3 Hz, 1H), 3.95 (dd, J = 1.5, 0.6 Hz, 2H), 3.79 (dd, J = 10.6,
4.0 Hz, 1H), 3.59 (dd, J = 10.6, 4.7 Hz, 1H), 1.54 (s, 9H), 1.47 (s, 3H), 1.41 (s, 3H).
13
C NMR
61
(151 MHz, Chloroform-d) δ 200.70, 168.87, 111.13, 81.84, 80.80, 77.87, 69.13, 68.52, 28.10,
27.04, 25.12.

Tert-butyl 2-(((4S,5R)-5-((E)-2-iodovinyl)-2,2-dimethyl-1,3-dioxolan-4-yl)methoxy)acetate
(3.3). Under inert atmosphere, CrCl
2
(1.50 g, 12.20 mmol) was added to a flame-dried flask that
was equipped with a stir bar and three-way adapter. Anhydrous THF (10 mL) was added via
syringe to the reaction flask. The solution was cooled to 0
o
C, followed by cannulation of
iodoform (1.92 g, 4.88 mmol) in 10 mL of THF. Next, the aldehyde (3.14, 0.298 g, 1.21 mmol)
was added via cannula, and the reaction was allowed to stir at 0
o
C for 3 hours, followed by 1
hour at room temperature. The reaction was quenched with water and then extracted with diethyl
ether (3 x 50 mL). The combined organic layers were washed with brine and dried over
anhydrous magnesium sulfate. The dried crude product was then filtered and concentrated in
vacuo, before purifying via flash column chromatography. The column was run at 10% ethyl
acetate: hexanes, and the eluted product was washed with an aqueous saturated solution of
sodium thiosulfate to remove any iodine side-product. The purified vinyl iodide (0.180 g, 37%)
was isolated.
1
H NMR (600 MHz, Chloroform-d) δ
H
6.56 (dd, J = 14.6, 6.7 Hz, 1H), 6.46 (dd, J
= 14.4, 0.9 Hz, 1H), 4.59 (td, J = 6.6, 1.0 Hz, 1H), 4.40 (td, J = 6.5, 5.3 Hz, 1H), 4.06-3.90 (m,
2H), 3.60-3.49 (m, 2H), 1.49 (s, 12H), 1.36 (s, 3H).
13
C NMR (151 MHz, Chloroform-d) δ
169.26, 141.20, 109.36, 81.76, 79.49, 79.14, 76.37, 70.24, 69.16, 28.12, 27.70, 25.17.
62

(2R, 3R)-3-(1,2-Dihydroxypropyl)-1,4-dioxaspiro[4,5]-decane-2-carboxaldehyde (3.15). A
solution of L-rhamnose (8.0 g, 43.92 mmol) and Drierite (2 g) in DMF (80 mL) were stirred at
0
o
C, under argon. The 1,1-dimethoxycyclohexane (13.4 mL, 88.09 mmol) was added via
syringe, followed by 40 mg (0.21 mmol) of p-toluenesulfonic acid. The reaction was stirred for
3 hours under argon at 0
o
C, then room temperature overnight. The reaction was quenched with 2
g of sodium carbonate, and allowed to stir for 1 hour. Then the mixture was filtered through
celite and concentrated in vacuo. The crude syrup was purified on silica by flash
chromatography, run at a gradient of 0% to 10% methanol: ethyl acetate. The purified product
was obtained in decent yield (7.51 g, 70%).
1
H NMR (500 MHz, Chloroform-d) δ
H
5.44 (s, 1H),
4.89 (dd, J = 5.9, 3.9 Hz, 1H), 4.62 (dd, J = 5.9, 0.8 Hz, 1H), 4.11-4.01 (m, 1H), 3.94 (dd, J =
7.3, 4.0 Hz, 1H), 1.70-1.51 (m, 10H), 1.34 (dd, J = 6.4, 0.8 Hz, 3H), 2.80-2.58 (m, 1H).

(2R, 3S)-α
2
-(1-Hydroxyethyl)-1,4-dioxaspiro[4,5]decane-2,3-dimethanol (3.16). Compound
3.16 was synthesized similarly to compound 3.12. No purification was performed; crude product
63
was used for the next step.
1
H NMR (500 MHz, Methanol-d
4
) δ
H
4.42 (dd, J = 6.9, 2.7 Hz, 1H),
4.22 (dd, J = 11.7, 5.8 Hz, 1H), 3.82-3.65 (m, 3H), 3.39 (dd, J = 7.8, 2.7 Hz, 1H), 1.74-1.56 (m,
9H), 1.42 (p, J = 6.0 Hz, 2H), 1.28-1.23 (m, 3H).
13
C NMR (126 MHz, Methanol-d
4
) δ 78.81,
76.79, 74.43, 69.48, 62.42, 37.89, 35.45, 26.41, 25.13, 24.82, 20.40.

Methyl 2-(((2S,3R)-3-(1,2-dihydroxypropyl)-1,4-dioxaspiro[4.5]decan-2-yl)methoxy)acetate
(3.17). In a round bottom flask, equipped with a three-way adapter and stir bar, the starting triol
(3.16, 0.75 g, 3.04 mmol) was added. Under argon, 20 mL of anhydrous THF was added to the
starting material, via syringe. The solution was cooled to 0
o
C, and then a slurry of t-BuOK (0.75
g, 6.70 mmol) in 8 mL THF was added via cannula. The reaction was stirred at 0
o
C for 30
minutes, after which methyl bromoacetate (0.35 mL, 3.68 mmol) was added by syringe over 10
minutes. The reaction was stirred at 0
o
C for 3 hours, and then room temperature overnight. The
mixture was concentrated in vacuo and purified on silica by flash chromatography (70% ethyl
acetate: hexanes). The purified product was collected along with the alternatively alkylated diols
(0.618 g, 63%), which are unreactive towards the next reaction. Mixture of three products:
1
H
NMR (500 MHz, Chloroform-d) δ
H
4.70 (s, 1H), 4.48-4.40 (m, 1H), 4.39-4.33 (m, 1H), 4.31 (d,
J = 0.8 Hz, 0H), 4.26-4.06 (m, 1H), 3.90 (dddd, J = 18.3, 9.8, 6.2, 0.8 Hz, 1H), 3.83-3.67 (m,
5H), 3.62 (dd, J = 6.0, 3.3 Hz, 1H), 1.71-1.48 (m, 10H), 1.30 (ddd, J = 6.4, 3.8, 0.8 Hz, 3H).
13
C
NMR (126 MHz, Chloroform-d) δ 170.57, 169.64, 167.71, 109.02, 108.98, 76.75, 75.79, 75.75,
64
75.46, 72.22, 70.88, 70.76, 70.05, 68.55, 68.31, 60.69, 52.40, 51.93, 36.98, 34.23, 25.08, 24.08,
23.70, 19.99, 19.93.

Methyl 2-(((2S,3S)-3-formyl-1,4-dioxaspiro[4.5]decan-2-yl)methoxy)acetate (3.18).
Compound 3.18 was synthesized similarly to compound 3.14. The pure product (0.90 g, 17%)
was obtained by column chromatography run at 50% ethyl acetate: hexanes.
1
H NMR (600
MHz, Chloroform-d) δ
H
9.60 (d, J = 2.3 Hz, 1H), 4.49 (dt, J = 7.7, 4.3 Hz, 1H), 4.34 (dd, J = 7.6,
2.3 Hz, 1H), 3.99 (d, J = 2.9 Hz, 2H), 3.70 (dd, J = 10.8, 4.0 Hz, 1H), 3.64 (s, 3H), 3.51 (dd, J =
10.8, 4.5 Hz, 1H), 1.62-1.46 (m, 10H).
13
C NMR (151 MHz, Chloroform-d) δ 200.95, 170.17,
128.28, 111.79, 80.41, 77.49, 68.80, 68.54, 51.78, 36.66, 34.47, 24.97, 23.95, 23.62.

Methyl 2-(((2S,3R)-3-((E)-2-iodovinyl)-1,4-dioxaspiro[4.5]decan-2-yl)methoxy)acetate (3.4).
Compound 3.4 was synthesized similarly to compound 3.3. The pure product (0.032 g, 49%)
was obtained by column chromatography run at 10% ethyl acetate: hexanes and the eluted
65
product was washed with an aqueous saturated solution of sodium thiosulfate to remove any
iodine side-product.
1
H NMR (500 MHz, Chloroform-d) δ
H
6.56 (ddd, J = 14.5, 6.5, 0.8 Hz,
1H), 6.49-6.47 (m, 1H), 4.59 (tt, J = 6.5, 0.9 Hz, 1H), 4.42-4.34 (m, 1H), 4.06 (td, J = 6.7, 0.8
Hz, 1H), 3.77 (d, J = 0.8 Hz, 3H), 3.75 (s, 1H), 3.55 (ddd, J = 5.7, 4.1, 0.8 Hz, 2H), 1.68-1.54
(m, 10H).
13
C NMR (126 MHz, Chloroform-d) δ
H
170.57, 141.31, 110.05, 79.46, 78.83, 75.96,
70.62, 68.71, 51.88, 37.55, 34.66, 25.03, 23.99, 23.65.

Tert-butyl-2-(((4S,5R)-2,2-dimethyl-5-(6-(trimethylsilyl)hexa-1,3-dien-5-yn-1-yl)-1,3-
dioxolan-4-yl)methoxy)acetate (3.20). A solution of the phosphonium salt (3.19, 0.195 g, 0.41
mmol) in 1 mL dry THF, under inert atmosphere, was stirred at -78
o
C. To the stirring mixture, n-
butyl lithium (0.18 mL, 0.32 mmol, 1.8 M in hexanes) was added and the reaction color changed
to orange. The temperature was raised to 0
o
C for 30 minutes, during which the reaction became
red. The temperature was lowered down to -78
o
C, and then a solution of the aldehyde (3.14,
0.050 g, 0.20 mmol) in THF (2 mL) was added via cannula. Next, the reaction was allowed to
warm to room temperature and stirred for 3.5 hours, before being quenched with ammonium
chloride. The product was extracted with diethyl ether (3 x 5 mL) and the combined organic
layers were dried over anhydrous magnesium sulfate, followed by filtration and concentration in
vacuo. The crude product (E:Z mixture 2:1 by
1
H NMR) was purified by silica gel column
66
chromatography, run at 8% ethyl acetate: hexanes, to afford 18 mg of the pure Z-isomer, 12 mg
of the pure E-isomer, and 16 mg of the Z/E-isomer mixture (0.046 g total, 57% yield). Each
purified isomer is characterized below.

tert-butyl 2-(((4S,5R)-2,2-dimethyl-5-((1Z,3E)-6-(trimethylsilyl)hexa-1,3-dien-5-yn-1-yl)-1,3-
dioxolan-4-yl)methoxy)acetate (3.20a).
1
H NMR (500 MHz, Chloroform-d) δ
H
6.88 (ddd, J =
15.6, 11.6, 1.2 Hz, 1H), 6.17 (t, J = 11.5 Hz, 1H), 5.67 (d, J = 15.1 Hz, 1H), 5.52 (t, 1H), 5.09
(ddd, J = 9.2, 6.7, 1.2 Hz, 1H), 4.43 (td, J = 6.9, 4.2 Hz, 1H), 3.98 (d, J = 7.6 Hz, 2H), 3.57-3.48
(m, 2H), 1.51 (s, 3H), 1.47 (s, 9H), 1.39 (s, 3H), 0.19 (d, J = 0.5 Hz, 9H).
13
C NMR (126 MHz,
Chloroform-d) δ 169.40, 136.53, 131.19, 128.84, 114.00, 109.27, 103.86, 98.79, 81.63, 73.08,
70.75, 69.21, 28.11, 27.88, 25.29, -0.15.

tert-butyl 2-(((4S,5R)-2,2-dimethyl-5-((1E,3E)-6-(trimethylsilyl)hexa-1,3-dien-5-yn-1-yl)-1,3-
dioxolan-4-yl)methoxy)acetate (3.20b).
1
H NMR (500 MHz, Chloroform-d) δ
H
6.63 (dd, J =
O
O O O
O
TMS
O
O O O
O
TMS
67
15.7, 10.9 Hz, 1H), 6.37-6.27 (m, 1H), 5.78 (ddd, J = 15.2, 7.3, 0.9 Hz, 1H), 5.64 (d, J = 15.7
Hz, 1H), 4.68 (t, J = 6.9 Hz, 1H), 4.42 (q, J = 6.1 Hz, 1H), 4.05-3.88 (m, 2H), 3.54-3.46 (m, 2H),
1.51 (s, 3H), 1.48 (d, J = 0.7 Hz, 9H), 1.37 (s, 3H), 0.19 (d, J = 0.8 Hz, 9H).
13
C NMR (126
MHz, Chloroform-d) δ 169.34, 141.51, 131.88, 131.09, 112.11, 109.18, 104.02, 97.85, 81.68,
70.72, 69.06, 28.13, 27.79, 25.23, -0.12.

(S,E)-((4-(3-bromophenyl)-1-(4-fluorophenoxy)but-3-en-2-yl)oxy)(tert-butyl)diphenylsilane
(3.22). Under inert atmosphere the vinyl iodide (3.1, 0.100 g, 0.183 mmol), 3-
bromophenylboronic acid (0.044 g, 0.219 mmol), K
2
CO
3
(0.076 g, 0.550 mmol), and Pd(PPh
3
)
4

(0.006 g, 0.005 mmol) were added to a screw-cap vial with a stirbar. Dioxane (3 mL) and
HPLC-grade water (1 mL) were also added to the vial, and the mixture was flushed with
nitrogen, before being sealed and heated at 90
o
C, overnight. The reaction was diluted with ethyl
acetate, washed with an aqueous saturated solution of sodium bicarbonate, and then brine. The
crude product was purified by column chromatography, run on silica gel at a gradient of 0% to
5% ethyl acetate: hexanes. The purified product (0.0267 g, 25%) was afforded as a white solid.
1
H NMR (600 MHz, Chloroform-d) δ
H
7.74-7.67 (m, 4H), 7.45-7.38 (m, 3H), 7.38-7.32 (m, 5H),
7.18-7.12 (m, 2H), 6.93-6.85 (m, 2H), 6.65-6.60 (m, 2H), 6.37 (dd, J = 15.9, 1.2 Hz, 1H), 6.22
(dd, J = 15.9, 6.3 Hz, 1H), 4.64 (tdd, J = 6.3, 5.5, 1.2 Hz, 1H), 3.96 (dd, J = 9.6, 6.2 Hz, 1H),
3.86 (dd, J = 9.6, 5.5 Hz, 1H), 1.11 (s, 9H).

68

(S,E)-tert-butyl((1-(4-fluorophenoxy)-4-(3-(4,4,5,5-tetramethyl-1,3,2-dioxaborolan-2-
yl)phenyl)but-3-en-2-yl)oxy)diphenylsilane (3.23). Under inert atmosphere, the aryl bromide
(3.22, 0.013 g, 0.0226 mmol), bis(pinacolato)diboron (0.007 g, 0.0276 mmol), potassium acetate
(0.008 g, 0.0815 mmol), and PdCl
2
(dppf) (0.001 g, 0.001 mmol) were added to a round bottom
flask with a stirbar. DMSO (0.75 mL) and dioxane (0.25 mL) were added and then the flask was
equipped with a septum with an argon balloon. The reaction was stirred at 80
o
C for 2 hours, and
then was quenched with ammonium chloride. The product was extracted with ethyl acetate (3 x
5 mL) and the combined organic layers were dried over anhydrous sodium sulfate, followed by
filtration and concentration in vacuo. The crude material was purified by preparative thin layer
chromatography, with a mobile phase of 10% ethyl acetate: hexanes. The purified product was
isolated in low yield (0.003 g, 21%).
1
H NMR (600 MHz, Chloroform-d) δ
H
7.76-7.64 (m, 7H),
7.45-7.28 (m, 7H), 6.91-6.83 (m, 2H), 6.63-6.58 (m, 2H), 6.49-6.43 (m, 1H), 6.27 (dd, J = 15.9,
6.5 Hz, 1H), 4.63 (tdd, J = 6.5, 5.3, 1.2 Hz, 1H), 3.94 (dd, J = 9.6, 6.4 Hz, 1H), 3.85 (dd, J = 9.6,
5.3 Hz, 1H), 1.36 (s, 12H), 1.10 (s, 9H).
13
C NMR (151 MHz, Chloroform-d) δ 156.36, 154.72,
136.03, 134.03, 132.79, 131.58, 129.64, 129.45, 128.90, 128.82, 127.88, 127.53, 126.34, 115.52,
83.86, 72.65, 65.85, 29.97, 29.70, 27.04, 24.87.

69

(S,E)-((4-(4-bromophenyl)-1-(4-fluorophenoxy)but-3-en-2-yl)oxy)(tert-butyl)diphenylsilane
(3.24). Compound 3.24 was synthesized similarly to compound 3.22, in 33% yield.
1
H NMR
(600 MHz, Chloroform-d) δ
H
7.75-7.66 (m, 5H), 7.46-7.29 (m, 9H), 7.12 (d, J = 8.4 Hz, 1H),
6.89 (dd, J = 9.1, 8.2 Hz, 1H), 6.68-6.58 (m, 2H), 6.40-6.30 (m, 1H), 6.21 (dd, J = 15.9, 6.4 Hz,
1H), 4.63 (tdd, J = 6.4, 5.4, 1.2 Hz, 1H), 3.96 (dd, J = 9.6, 6.2 Hz, 1H), 3.87 (dd, J = 9.6, 5.4 Hz,
1H), 1.10 (s, 9H).
13
C NMR (151 MHz, Chloroform-d) δ 157.99, 156.41, 154.68, 135.98,
135.93, 135.61, 134.78, 133.78, 133.58, 131.55, 130.47, 129.71, 129.62, 128.03, 127.70, 127.54,
121.38, 115.73, 115.58, 115.50, 115.45, 72.47, 72.42, 29.70, 29.70, 27.00, 26.99, 26.55, 19.40.

(S,E)-tert-butyl((1-(4-fluorophenoxy)-4-(4-(4,4,5,5-tetramethyl-1,3,2-dioxaborolan-2-
yl)phenyl)but-3-en-2-yl)oxy)diphenylsilane (3.25). Compound 3.25 was synthesized similarly
to compound 3.23, in 19% yield.
1
H NMR (400 MHz, Chloroform-d) δ
H
7.77-7.64 (m, 7H),
7.59-7.50 (m, 1H), 7.46-7.28 (m, 8H), 6.93-6.84 (m, 2H), 6.67-6.59 (m, 2H), 6.46-6.38 (m, 1H),
6.27 (dd, J = 15.9, 6.4 Hz, 1H), 4.64 (tdd, J = 6.4, 5.3, 1.1 Hz, 1H), 3.96 (dd, J = 9.6, 6.3 Hz,
1H), 3.87 (dd, J = 9.6, 5.3 Hz, 1H), 1.35 (s, 12H), 1.09 (s, 9H).

70
3.5 Chapter 3 References

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12
Fiorucci, S.; Wallace, J. L.; Mencarelli, A.; Distrutti, E.; Rizzo, G.; Farneti, S.; Morelli, A.;
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Clish, C. B.; O’Brien, J. A.; Gronert, K.; Stahl, G. L.; Petasis, N. A.; Serhan, C. N. Proc. Natl.
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71

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Takai, K.; Nitta, K.; Utimoto, K. J. Am. Chem. Soc. 1986, 108, 7408.

20
Ramirez, F.; Desai, N. B.; McKelvie, N. J. Am. Chem. Soc. 1962, 84, 1745.

21
Corey, E. J.; Fuchs, P. L. Tetrahedron Lett. 1972, 36, 3769.

22
Barbat, J.; Gelas, J.; Horton, D. Carbohydrate Research 1983, 116, 312.

23
Munier, P.; Krusinski, A.; Picq, D.; Anker, D. Tetrahedron 1995, 51, 1229.

24
Rerat, V.; Dive, G.; Cordi, A. A.; Tucker, G. C.; Bareille, R.; Amedee, J.; Bordenave, L.;
Marchand-Brynaert, J. J. Med. Chem. 2009, 52, 7029.

25
Miyaura, N.; Yamada, K.; Suzuki, A. Tetrahedron Lett. 1979, 36, 3437.

26
Miyaura, N.; Suzuki, A. J. C. S. Chem. Comm. 1979, 19, 866.

27
Ishiyama, T.; Murata, M.; Miyaura, M. J. Org. Chem. 1995, 60, 7508.

72
Chapter 4. Synthesis of Potential Modulators of GRP78
4.1 Introduction
The endoplasmic reticulum (ER) is responsible for the synthesis, folding, and transport of
proteins in the cell. Under a state of stress, such as glucose deprivation, the endoplasmic
reticulum slows down its protein folding and this results in an accumulation of unfolded proteins
in the ER. The excess of unfolded and misfolded proteins in the ER triggers the aptly-named
“unfolded protein response” (UPR).
1,

2, 3
The UPR begins with signaling chaperone proteins to
fold the unfolded proteins and the degradation of misfolded proteins in an effort to relieve the
stressful environment.
2, 3
If the stress of the ER cannot be ameliorated, then the UPR signals
apoptosis and the cell dies.
2, 3
ER homeostasis and the UPR pathway are vital to mammalian
development and health.
4
ER stress has been linked to several diseases, such as Alzheimer’s and
Parkinson’s disease, cancer, and diabetes.
5,

6

78,000-Dalton glucose-regulated protein (GRP78) is a chaperone protein in the ER that
has a crucial role in modulating the UPR. GRP78 is bound to three different transmembrane
proteins (kinases IRE1 and PERK, and transcription factor ATF6) when the ER is in a state of
homeostasis.
7
GRP78 prevents these transmembrane protein sensors from aggregating and this
keeps them inactive.
1
Under ER stress, GRP78 dissociates from each of the transmembrane
proteins and this activates them by allowing them to aggregate, ultimately resulting in a decrease
in protein translation and an increase in protein folding inside the ER.
3,

7

Tumor cells metabolize glucose at a much faster rate than normal cells, and this causes a
state of ER stress and UPR activation that leads to GRP78 accumulation and over expression in
cancer.
2
GRP78 has been found to prevent several actions of apoptosis, causing cancer cell
73
survival and protection.
8
GRP78 binds Ca
2+
, removing it from the ER and preventing the Ca
2+

from entering the cytosol and signaling apoptosis.
9,

10
Another way that GRP78 blocks cell death
is by interacting with enzymes that regulate apoptosis, such as BIK, BAX, caspase-7 and
caspase-12.
2,

11
Importantly, apoptotic activity can be restored to the cell by inhibiting or
knocking down GRP78 activity. Knocking down GRP78 also restores drug sensitivity to drug
resistant tumor cells, specifically in malignant glioma.
6, 12
Adjuvant cancer therapy could
possibly benefit from this re-sensitization of chemoresistant malignant tumors by administering a
treatment that suppresses GRP78 in addition to the primary cancer treatment. Also, breast and
prostate cancer progression, reoccurance, and survival can be predicted from the over-expression
of GRP78.
13, 14, 15, 16
Therefore, GRP78 expression could be used as a marker in cancer
diagnosis, as well as in determining a course of treatment.
Interestingly, it was recently discovered that GRP78 is not confined to the ER in cancer
cells, but rather it migrates to the cell surface.
17,

18
Although the exact mechanism of how
GRP78 travels to the outer plasma membrane is not completely elucidated, it has been
established that cell surface GRP78 is a receptor for cancer signaling proteins, specifically α2-
macroglobulin, Kringle 5, and GRP78 binding peptide (see Figure 4.1).
19,

20, 21
Angiogenesis
inhibitor Kringle 5 binds to cell surface GRP78 and blocks it, restoring the apoptotic activity to
the cell.
22
Activated α2-macroglobulin can also bind to cell surface GRP78 as a receptor, which
is believed to play a role in prostate cancer metastasis.
19, 23
GRP78 binding peptide, also referred
to as ADoPep1, binds to cell surface GRP78 and causes apoptosis in astrocytes and neurons.
24

Due to the obvious significance of cell surface GRP78 in many cancer-signaling pathways, and
its presence at the cell surface only in cancer cells, GRP78 is an attractive target for possible
anticancer therapeutics.
74
Various natural compounds have also shown anticancer activity due to their inhibition of
GRP78.
16
Figure 4.1 shows the known natural products that have an inhibitory effect on GRP78
and where the natural products are derived from. Notably, the structures of GRP78 inhibitors
vary greatly, from antioxidant small molecules such as epigallocatechin gallate (EGCG) found in
green tea, all the way up to the protein interleukin 24 (IL-24). The inhibitors of GRP78 also vary
greatly in specificity. EGCG is known for broad anticancer activity by acting on several
differerent proteins, whereas protein AB
5
subtilase toxin from E. coli was discovered to cause
apoptosis in cancer cells by cleaving one specific amino acid in GRP78.
25
While small
molecules have the advantage of greater stability and ease of synthesis, compared to proteins,
they are generally less specific because they can easily bind in several different active sites.

Figure 4.1- Stress-induced GRP78 activity within a tumor cell.
16
GRP78 migrates to the surface
of the cell, and this may critical to cancer metastasis.
neuroblastoma that requires further investigation (13, 14). Auto-
antibodies against GRP78 at high levels in prostate cancer patients
have also been reported, associating with aggressive tumor
behavior (29, 30). Furthermore, expression of GRP78 was recently
examined retrospectively in prostate cancer patients during the
development of castration resistance (31). GRP78 is strongly up-
regulated during the transition from localized prostate cancer to
metastatic castrationand may serve asa novelprognosticindicator
of recurrence in untreated patients with localized tumor.
In the management of breast cancer patients, there are at
present only two biomarkers that are used to predict potential
benefits of adjuvant therapy for the disease, hormone receptor
status, and Her2/neu status. The utility of these biomarkers to
eliminate ineffective treatment cannot be underestimated. For
women with hormone receptor–negative tumors, adjuvant hor-
monal therapy will not reduce the risk of recurrence. Therefore,
such women may be spared the toxicities of such agents. Similarly,
patients whose tumors do not overexpress Her2/neu may be
spared treatment with Herceptin. Unfortunately, similar tests that
would predict benefit from adjuvant systemic chemotherapy
agents do not exist. Such tests would be useful in avoiding the
toxicity associated with chemotherapy in patients who would not
benefit from treatment with these agents. Based on preclinical
studies strongly suggesting that GRP78-positive tumors may be
resistant to topoisomerase inhibitors (5, 16–20), a retrospective
study was conducted to evaluate the value of GRP78 as a
biomarker for treatment response. The study revealed that two-
thirds of breast cancer patients show moderate to high levels of
GRP78 in biopsies before treatment, and that in patients who
received adjuvant systemic chemotherapy with Adriamycin-based
regimens, GRP78 positivity indicated a higher risk of recurrence
(32). Thus, upon validation, GRP78 positivity might identify
patients who could be spared the toxicities of Adriamycin-based
adjuvant chemotherapy. Another observation that warrants addi-
tional investigation is whether GRP78 positivity might also identify
patients who are more likely to benefit from treatment with
Adriamycin followed by taxanes (32). In preclinical studies with
estrogen-positive human breast cancer cells, GRP78 confers
resistance to estrogen starvation–induced apoptosis through
suppressing the activity of BIK (21). Thus, GRP78 level may also
Figure 1. ER stress induction of GRP78in the tumor microenvironment. Both intrinsic and extrinsic factors lead tothe up-regulation ofGRP78 (also referredto asBiP)
and its cell surface expression in tumor cells. Through inhibition of apoptosis, GRP78 facilitates tumor progression, immune resistance, metastasis, and drug
resistance. Dormant tumor cells, as well as quiescent tumor endothelial cells, also rely on GRP78 to escape chemotherapy. Anticancer compounds that either inhibit
the stress induction of GRP78 or suppress its catalytic function have been identified from multiple sources. When used in combination therapy, they should
enhance drug efficacy, lower resistance and assist in eradicating residual tumor. GRP78 has also been identified as a cell surface receptor for Kringle 5 of human
plasminogenandtheactivatedformofthea2-macroglobulin.BecausecellsurfaceGRP78isnotdetectedinnormalorgans,itcanserveasaconduitforcancer-specific
delivery of cytotoxic agents via GRP78 binding peptides. Autoantibody levels against GRP78 in patient serum, as well as expression levels of GRP78 in biopsies,
may represent novel biomarkers in stratifying patients for tumor behavior and responsiveness to treatment. Abbreviations: N, nucleus; ER, endoplasmic reticulum;
C, cytoplasm.
Cancer Research
Cancer Res 2007; 67: (8). April 15, 2007 3498 www.aacrjournals.org
Research.
on August 6, 2013. © 2007 American Association for Cancer cancerres.aacrjournals.org Downloaded from
75
4.2 Results and Discussion
4.2.1 Design of a Small Library of Compounds for Testing GRP78 Activity
Given the great significance of GRP78 overexpression in cancer and its essential role in
cell survival, drug-resistance, and possibly cancer metastasis, we wanted to synthesize small
molecules that could modulate GRP78. Although certain peptides have interesting and impactful
biological activity, proteases and other enzymes easily break them down in vivo. We wanted to
combine the specificity of a protein with the stability and ease of delivery of a small molecule,
thus we turned our attention to peptidomimetics. Peptidomimetics are peptide-like compounds
that have some structural element (such as an unnatural amino acid) that adds stability or
biological activity to the compound. Peptidomimetics can be made completely from small
molecule building block, such as amino acids, or pre-exhisting peptides can be chemically
altered for a specific purpose. Importantly, peptidomimetics can be designed to target a specific
protein-protein interaction or protein structural motif.
Our particular interest was in synthesizing a specific class of peptidomimetic molecules
called “reverse-turn mimetics”. Reverse-turn mimetics are peptidomimetics that imitate the rigid
U-shaped conformation of a β-turn found in naturally-occuring peptides.
26
The β-turn of native
peptides is held together through hydrogen bonding interactions between amino acids. A
reverse-turn motif can have enhanced stability compared with a β-turn by replacing the native
hydrogen bonds with covalent bonds. Reverse-turn mimetic small molecules have great
potential in the area of cancer signaling pathways, as shown by McMillan and Kahn when they
used a tetrahydropyrazinopyrimadinedione to block a specific protein-protein interaction of the
Wnt-β-catenin signaling pathway, a pathway critical to cancer development.
27,

28

76
A new class of reverse-turn peptidomimetics called hexahydropyrazinotriazinediones has
recently been synthesized and is being investigated for potential medicinal properties. Kim and
coworkers found that micromolar quantities of some hexahydropyrazinotriazinediones enhance
cardiogenesis in human stem cells.
29
The most active compound of the screening was CW209E,
a reverse-turn mimetic with the same underlying ring structure as shown in Figure 4.2. The
authors investigated whether CW209E was acting on the gene expression and/or protein-protein
interactions.
29
While Kim et. al. discovered that the active reverse-turn mimetic was
upregulating specific cardiomyogenic genes, they were unable to ascertain if the molecule was
also interfering with a protein-protein interaction.
29
However, it is well known that protein-
protein interactions are complex, and not all of them are fully elucidated, so it is probable that
CW209E could also be inferring with a protein-protein interaction, but further studies are needed
to determine this.
29
Given these useful findings, we sought to design novel
hexahydropyrazinotriazinediones which could have significant effects on modulating GRP78,
either through protein-protein interactions or through gene expression.
Figure 4.2 shows the general template of the design of the desired compounds to
synthesize for GRP78 activity testing. Note that each one of the substituted functional groups
(R, R’, or R”) could either have an aromatic ring, or an unactive “blocking group” of a methyl or
hydrogen. These changes to the structure would allow us to investigate the structure-activity-
relationship (SAR) between aromatic side chains and the active site of the GRP78 protein.

77

Figure 4.2- The design of novel hexahydropyrazinotriazinedione β-turn peptidomimetics for the
modulation of GRP78.

Based on the structure shown in Figure 4.2, we designed three novel
hexahydropyrazinotriazinediones (See Figure 4.3) which could serve as potential GRP78
modulators, 4.1, 4.2, and 4.3. We wanted to investigate whether or not an aromatic ring
placement had any marked effect on the biological activity of the molecule. An aromatic ring is
a large functional group, and so if it adds too much bulk to the molecule, it could inhibit a
possible desirable interaction with GRP78. On the other hand, an aromatic ring in the correct
placement and orientation on the molecule could aid in valuable pi-interactions with GRP78.
Notably, we are not altering the fundamental pyrazinotriazine fused bi-cyclic moiety whatsoever.
This allows us a two-fold advantage: one, we will not be introducing several variables of
structural change which cause biological data to be difficult to interpret, and two, we can apply
the same general synthetic strategy towards different molecules.
N
N
N
N
O NH
O
O
R R"
R'
Aromatic group
or Hydrogen
Aromatic group
or Hydrogen
Aromatic group
or Methyl
78

Figure 4.3- Structures of novel final compounds for potential GRP78 modulation synthesized
herein.

4.2.2 Retrosynthesis of Potential Modulators of GRP78
Scheme 4.1 shows the general retrosynthesis for the reverse-turn mimetic potential
modulators of GRP78. The fused bi-cyclic final compounds can be synthesized from the
cyclization of a peptide-like backbone structure (4.4). The stereochemistry of R’ is determined
from the amino acid chosen as a starting material in the beginning of the synthesis.

Scheme 4.1- The retrosynthesis of hexahydropyrazinotriazinedione final compounds from a
peptide-like backbone structure.

N
N
N
NH
O NH
O
O
N
N
N
N
O NH
O
O
S
N
N
N
N
O NH
O
O
S
4.1 4.3 4.2
N
N
N
N
R
O NH
O
O
N
H
N
H
O
N
N
H
O
N
O
O
O
4.4
R"
R'
R R' R"
R = CH3 or
R' = H or
R" = H or S
79
The overall retrosynthesis of the peptide-like backbone is shown in Scheme 4.2. The
peptide-like backbone (4.4) is derived from two different building blocks, an amine (4.6) and a
carboxylic acid (4.5) that are bonded using a peptide coupling reaction. The carboxylic acid
(shown in blue, 4.5) is synthesized from benzyl amine and a boc-protected hydrazine.
Compound 4.6 is made from coupling an amino acid with an aminoacetaldehyde diethyl acetal.
The diethyl acetal group is removed during the final cyclization of the product, but serves as a
useful protecting group throughout the synthesis.

Scheme 4.2- The retrosynthesis of 4.4, the general peptide-like backbone, derived from building
blocks which are synthesized from readily available commercial starting materials, such as
amino acids and benzyl amine.

N
H
N
H
O
N
O
OH
H
2
N
N
O
O
O
NH
2
H
2
N
N
Boc
H
2
N
O
OH HN
O
O
4.5 4.6
R
R' R"
R
R' R"
N
H
N
H
O
N
N
H
O
N
O
O
O
4.4
R R' R"
80
4.2.3 Synthesis of Potential GRP78 Modulator Building Blocks
In order to synthesis title compounds 4.1, 4.2, and 4.3, we first needed to synthesize both
carboxylic acid building blocks and amine building blocks that could be reacted together and
later cyclized. The two carboxylic acids we wanted to synthesize were 4.7 and 4.8, shown in
Figure 4.4, and three amine building blocks, 4.9, 4.10 and 4.11. The two carboxylic acid
building blocks vary only by the functionality on the hydrazine (methyl versus benzyl), however
they required a slightly different synthetic method, as is shown later. The three amines vary by
either containing a thiophene and/or a benzyl moiety.

Figure 4.4- Essential carboxylic acid building blocks and amine building blocks for the synthesis
of the the title compounds.
N
H
N
H
O
N
OH
O
N
H
N
H
O
N
OH O
H
2
N
N
O
S
O
O
H
2
N
N
O
S
O
O
H
2
N
N
H
O
O
O
4.7
4.8
4.9
4.10
4.11
81
Carboxylic acid 4.7 was synthesized in four overall steps, each of which gave at least a
60% yield (Scheme 4.3). The synthesis begins with commercially available starting materials,
benzyl amine and 1-boc-1-methylhydrazine, being coupled with 1,1’-carbonyldiimidazole (CDI)
at 0
o
C to yield the white solid product (4.12), which was recrystallized. Next, 4.12 was
deprotected using 4M HCl in dioxane at room temperature, and the reaction was stirred
overnight. Product 4.13 was a white solid that was purified via recrystallization. An SN
2

reaction was then used to transform 4.13 to 4.14, using t-butyl bromoacetate and K
2
CO
3
as a
base, and heating the reaction at 80
o
C overnight. Finally, the t-butyl group was removed from the
carboxylic ester (4.14) using 4M HCl in dioxane at 0
o
C, to produce the desired carboxylic acid
(4.7) building block in 74% yield.

Scheme 4.3- The synthesis of carboxylic acid 4.7 in four overall steps.
N
H
N
H
O
N O
O
NH
2 H
2
N
N O
O
+
a
b
N
H
N
H
O
NH
N
H
N
H
O
N
O
O
c
d
N
H
N
H
O
N
OH
O
4.12
4.13
4.14
4.7
Reagents and conditions: (a) 1,1'-carbonyldiimidazole, triethyl amine, DMF, 0
o
C to rt,overnight, 78%
(b) 4M HCl in dioxane, rt, overnight, 60% (c) t-butyl bromoacetate, K
2
CO
3
, toluene, DMF, 80
o
C,
overnight, 75% (d) 4M HCl in dioxane, 0
o
C to rt, overnight, 74%
82
The same synthetic method that was successful for 4.7 was attempted for the similar
carboxylic acid building block 4.8. The synthesis of 4.8 begins with a boc-protection of the
commercially available benzylhydrazine hydrochloride to yield 4.15 (Scheme 4.4). With
hydrazine 4.15 in hand, a CDI coupling reaction was performed to yield 4.16 in 44% yield.
Although the conditions were the same for this reaction as the CDI coupling shown in Scheme
4.3, the bulkiness of the benzyl group probably cause the reduction in product yielded. Next,
4.17 was synthesized by deprotecting 4.16 using TFA and stirring at room temperature for 5
hours, and was used crude for the next step. It was found that while most of the same reaction
conditions from Scheme 4.3 were successful for Scheme 4.4, the SN
2
reaction proved to be
problematic. The reason that the SN
2
reaction was very successful in the synthesis of 4.7 but not
4.8 was likely due to the small size of the methyl group on 4.7 compared to the bulkiness of the
benzyl group on 4.8. To alleviate this problem, we decided to search for an alternative reaction
that would not be as sensitive to the bulkiness of the benzyl group on 4.17. Luckily, a reductive
amination was found to be successful in producing the final product 4.8 in 45% yield. While the
yield is not extremely high, it was more than expected for using crude starting material. The
reductive amination allowed us to use glyoxylic acid monohydrate, meaning that there was no
need for a protection or deprotection of the final carboxylic acid group, saving us time and steps.

83

Scheme 4.4- The synthesis of carboxylic acid 4.8 utilizing a reductive amination in place of the
SN
2
reaction used for the synthesis of 4.7.

Next, the amine building blocks (4.9, 4.10, and 4.11) were synthesized as shown in
Schemes 4.5, 4.6, and 4.7. The amine 4.18 was synthesized via a reductive amination between
the thiophene-2-aldehyde and aminoacetaldehyde diethyl acetal, heated at 80
o
C, neat, and then
the reaction was cooled to room temperature and ethanol and sodium borohydride were added
and the reaction was stirred overnight. Amine 4.18 was used to synthesize both 4.9 and 4.10. To
synthesize 4.9, piece 4.18 underwent a peptide coupling reaction with FMOC-protected glycine
using the coupling reagent HATU (1-[Bis(dimethylamino)methylene]-1H-1,2,3-triazolo[4,5-
b]pyridinium-3-oxid hexafluorophosphate) (see Scheme 4.5). The peptide coupling reaction was
very successful, giving product 4.19 in 99% yield. The final piece 4.9 was produced in modest
yield by deprotecting the FMOC functional group on 4.19 using diethyl amine (DEA) and mild
conditions.
H
2
N
N O
O
H
2
N
NH
a
4.15
N
H
N
H
O
N O
O
4.16
b
c
N
H
N
H
O
NH
4.17
d
N
H
N
H
O
N
4.8
O OH
Reagents and conditions: (a) di-tert-butyl-dicarbonate, THF, sodium bicarbonate solution added over 2 hours
at rt, 89% (b) 1,1'-carbonyldiimidazole, benzyl amine, triethyl amine, DMF, 0
o
C to rt,overnight, 44% (c) TFA,
DCM, 5 hours at rt (d) glyoxylic acid monoydrate, MeOH, NaBH
3
CN, stirred for 30 min. at rt, 45% yield
84

Scheme 4.5- The synthesis of amine building block 4.9 in three steps.
Amine 4.10 was synthesized similarly to 4.9, beginning with piece 4.18 and peptide
coupling it to FMOC-protected phenylalanine (Scheme 4.6). The same method of peptide
coupling reaction was consistently successful for several different pieces synthesized herein.
Again, the FMOC protecting group was deprotected using DEA, but this time the reaction was
much higher yielding than previously.

Scheme 4.6- The synthesis of 4.10 using a peptide coupling reaction and a deprotection.
HN
S
O
O
a
NH
2
O
O
+
S
H O
b
N
H
O
O
O
S
N
O
O
c
4.18
4.19
H
2
N
O
O
O
S
N
4.9
Reagents and conditions: (a) 80
o
C, Neat for 20 min., then cooled to rt, added ethanol and NaBH
4
, stirred overnight, 71%
(b) FMOC-Gly-OH, DMF, Hunig's base, HATU, stir at rt for 30 min. then the amine was added and stirred at rt overnight,
99% (c) Diethyl amine, DCM, stir at rt overnight, 53%.
N
H
N O
S
O
O
O
O
HN
S
O
O
4.18
a
4.20
b
H
2
N
N O
S
O
O
4.10
Reagents and conditions: (a) FMOC-Phe-OH, DMF, Hunig's base, HATU, stir at rt for 30 min. then the amine was added
and stirred at rt overnight, 86% (b) Diethyl amine, DCM, stir at rt overnight, 89%.
85
Synthesis of the final amine building block (4.11) is shown in Scheme 4.7. Again, the
robust peptide coupling reaction was successful for the production of the peptide bond between
aminoacetaldehyde diethyl acetal and FMOC-Phe-OH. 4.21 was produced in 91% yield. The
FMOC deprotection reaction of 4.21 to produce 4.11 using DEA was low yielding, but gave
enough product to move ahead with the synthesis.

Scheme 4.7- The synthesis of amine 4.11 in two steps.

4.2.4 Assembly of Building Blocks into Final Products
Once all of the carboxylic acid and amine building blocks were in hand, we needed to
assemble the pieces together to make the linear backbone that would later be cyclized to the final
products. Again, a peptide coupling reaction using HATU as the coupling reagent was utilized
for all of the couplings shown in Schemes 4.8, 4.9, and 4.10. Products 4.22, 4.23, and 4.24 were
used crude without formal purification or characterization. This was because it was found that
H
2
N
O
O
a
H
N
N
H
O
O
O
O
O
H
N
O
O
O
OH
+
b
H
2
N
N
H
O
O
O
4.11
4.21
Reagents and conditions: (a) DMF, Hunig's base, HATU, stir at rt for 30 min. then the amine was added and stirred
at rt overnight, 91% (b) Diethyl amine, DCM, stir at rt overnight, 27%
86
the peptide coupling reaction is realiable in producing the desired product, and the by-products
of the reaction were found to be very difficult to separate from the product via column
chromatography. Also, once the crude linear peptide-like products have been cyclized they
would be much easier to purify.
Scheme 4.8 shows the peptide coupling reaction between carboxylic acid 4.8 and amine
4.11. The reaction required only mild conditions and stirring overnight at room temperature. As
stated previously, crude product 4.22 was used without further purification. Because –NH is a
good hydrogen bond donor and C=O is a good hydrogen bond acceptor, and these functional
groups are both abundant in this molecule, impurities that can hydrogen bond could easily be
trapped and make the purification of 4.22 difficult. The presence of the three bulky benzyl
groups on 4.22 could be the reason for a more problematic synthesis overall. However, as long
as each reaction was able to yield some amount of product, the synthesis was pushed forward
without attempts to improve the yield.

Scheme 4.8- Peptide coupling to produce 4.22, the precursor of final product 4.1.
H
2
N
N
H
O
O
O
4.11
N
H
N
H
O
N
4.8
O OH
+
a
N
H
N
H
O
N
HN
O
NH O
O
O
4.22
Reagents and conditions: (a) DMF, Hunig's base, HATU, stir at rt for 30 min. then the amine was added and stirred
at rt overnight
87
The synthesis of 4.23 is shown in Scheme 4.9. The peptide coupling reaction used
previously was again successfully utilitzed for the synthesis of 4.23. Compound 4.23 was used
crude without purification the same as compound 4.22. Below shows the same type of peptide
coupling between the carboxylic acid 4.7 and the amine 4.10 to produce molecule 4.24. Again,
4.24 was used crude without a need for purification.

Scheme 4.9- The synthesis of 4.23 using HATU as a peptide coupling reagent.

Scheme 4.10- The synthesis of backbone 4.24 to be later cyclized into 4.3.

N
H
N
H
O
N
4.8
O OH
+
H
2
N
O
O
O
S
N
4.9
a
N
H
N
H
O
N
HN
O
N O
S
O
O
4.23
Reagents and conditions: (a) DMF, Hunig's base, HATU, stir at rt for 30 min. then the amine was added and stirred
at rt overnight
N
H
N
H
O
N
OH
O
4.7
+
H
2
N
N O
S
O
O
4.10
a
N
H
N
H
O
N
N
H
O
N O
S
O
O
4.24
Reagents and conditions: (a) DMF, Hunig's base, HATU, stir at rt for 30 min. then the amine was added and stirred
at rt overnight
88
Schemes 4.11, 4.12, and 4.13 show the last step in the synthesis of title compounds 4.1,
4.2 and 4.3, respectively. Adding pure formic acid to the neat starting compound (either 4.22,
4.23 or 4.24) and allowing the reaction to stir overnight at room temperature allowed us to
cyclize the straight chain molecules into the bi-cyclic hexahydropyrazinotriazinedione final
products. Although these cyclization reactions were low yielding, they produced a much more
easily separable product compared to the starting material. Also, the low yield could be
attributed partially to the presence of impurities adding mass to the starting material. Overall,
the three final compounds were synthesized successfully and are currently being tested for
GRP78 structure-activity-relationship studies in cancer cell lines.

Scheme 4.11- The final step in the synthesis of final product 4.1.

N
H
N
H
O
N
HN
O
NH O
O
O
4.22
a
N
N
N
NH
O NH
O
O
4.1
Reagents and conditions: (a) formic acid, rt, overnight, 8%
89

Scheme 4.12- The final step in the synthesis of final product 4.2.

Scheme 4.13- The final step in the synthesis of final product 4.3.

N
H
N
H
O
N
HN
O
N O
S
O
O
4.23
a
N
N
N
N
O NH
O
O
S
4.2
Reagents and conditions: (a) formic acid, rt, overnight, 7%
N
H
N
H
O
N
N
H
O
N O
S
O
O
4.24
a
N
N
N
N
O NH
O
O
S
4.3
Reagents and conditions: (a) formic acid, rt, overnight, 10%
90
4.3 Conclusion
Novel hexahydropyrazinotriazinediones were successfully designed based on a reverse-
turn mimetic structure for potential GRP78 modulation. Alterations to the position of the
aromatic functional groups placed on the fused bi-cyclic ring structure were carefully chosen as a
means of investigating the structure-activity relationship between the final compounds and
GRP78. The potential GRP78 modulators were synthesized by assembling builing blocks that
could easily be combined using a peptide coupling reaction. Once the pieces were assembled,
the backbone was cyclized into the final final compound. Further studies are ongoing to
investigate the biological activity of each product and its interactions with GRP78.

4.4 Experimental
All reactions, unless otherwise noted, were carried out with commercially available
starting materials and solvents, which were used without further purification. The majority of
the chemicals were purchased from Sigma Aldrich, GFS chemicals, TCI America, and VWR
International. All glassware and reaction vessels used were standard and commercially
available. Reactions performed under “inert atmosphere” implies the reaction was run in a round
bottom flask affixed with a three-way adapter and an argon balloon, or the flask was affixed with
a septum which was punctured with a needle affixed to an argon balloon.
1
H and
13
C NMR
spectra were collected using a Varian Mercury 400, Varian 400-MR, Varian 500, or Varian 600
NMR spectrometers, using residual
1
H and
13
C peaks of deuterated solvents as internal standards.
Manual flash column chromatography was run with silica gel purchased from Sorbent
Technologies (60 Å, 40-63 µm). A Biotage Isolera One was used for automated flash column
91
chromatography, with standard Biotage silica cartridges (60 Å) for purification. LC-MS analysis
was performed on an Agilent Technologies 1200 infinity series instrument.

Procedure

tert-butyl 2-(benzylcarbamoyl)-1-methylhydrazinecarboxylate (4.12). To a round bottom
flask with a stirbar was added 1,1’-carbonyldiimidazole (1.22 g, 7.52 mmol) after which the flask
was equipped with a septum and argon balloon. DMF (7 mL) was added via syringe and the
solution was cooled to 0
o
C. Next, 1-boc-1-methylhydrazine (1.0 mL, 6.74 mmol), benzylamine
(0.75 mL, 6.87 mmol), and triethyl amine (1.05 mL, 7.52 mmol) were added via syringe and the
reaction was warmed to room temperature and then allowed to stir for 48 hours. The crude
product was recrystallized with hot ethyl acetate and hexane to yield pure white crystals (1.49 g,
78% yield).
1
H NMR (500 MHz, Chloroform-d) δ
H
7.37-7.26 (m, 5H), 6.16 (s, 1H), 5.34 (t, J =
6.1 Hz, 1H), 4.46 (d, J = 5.9 Hz, 2H), 3.12 (d, J = 0.7 Hz, 3H), 1.43 (s, 9H).
13
C NMR (126
MHz, Chloroform-d) δ 157.62, 155.89, 138.76, 128.65, 127.49, 109.98, 82.24, 43.88, 38.59,
28.09.

N
H
N
H
O
N O
O
92

N-benzyl-2-methylhydrazinecarboxamide (4.13). The solid starting material (4.12, 0.97 g,
3.47 mmol) was dissolved in 26 mL of dioxane and stirred at room temperature, then 5.2 mL of
HCl was added dropwise to reach a 4M concentration. The reaction was stirred at room
temperature overnight. The reaction mixture was then concentrated in vacuo, and then quenched
at 0
o
C with saturated sodium bicarbonate until the pH was approximately 8. The solution was
extracted with ethyl acetate (3 x 50 mL) and the combined organic layers were dried over sodium
sulfate. The solution was filtered and concentrated in vacuo, the product was recrystallized with
ethyl actate and hexane to yield 0.375 g (60% yield) of white crystals.
1
H NMR (500 MHz,
Chloroform-d) δ
H
7.37-7.27 (m, 5H), 6.42 (s, 1H), 5.54 (s, 1H), 4.45 (d, J = 6.1 Hz, 2H), 3.54 (s,
1H), 2.62-2.58 (m, 3H).

tert-butyl 2-(2-(benzylcarbamoyl)-1-methylhydrazinyl)acetate (4.14). To the hydrazine
(4.13, 0.300 g, 1.67 mmol) in a round bottom flask with a stirbar was added toluene (32 mL),
DMF (4 mL), and potassium carbonate (1.16 g, 8.39 mmol). The reaction was refluxed at 80
o
C
for 1 hour, cooled to room temperature, and then t-butyl bromoacetate (0.27 mL, 1.83 mmol) was
added and the reaction was refluxed at 80
o
C overnight. The reaction was then cooled to room
temperature, filtered, concentrated, and diluted with 25 mL of ethyl acetate. The solution was
N
H
N
H
O
NH
N
H
N
H
O
N
O
O
93
washed with brine (3 x 25 mL), dried over sodium sulfate, filtered, and then concentrated in
vacuo. The solid was recrystallized with hexane to yield 0.366 g of white solid in 75% yield.
1
H
NMR (400 MHz, Chloroform-d) δ
H
7.40-7.26 (m, 5H), 6.41 (t, J = 5.3 Hz, 1H), 6.19 (s, 1H),
4.45 (d, J = 6.0 Hz, 2H), 2.70 (s, 3H), 1.47 (s, 9H).

2-(2-(benzylcarbamoyl)-1-methylhydrazinyl)acetic acid (4.7). The protected hydrazine
starting material (4.14, 0.360 g, 1.23 mmol) was dissolved in 1.5 mL dioxane and then cooled to
0
o
C. Next, 0.5 mL of HCl was added dropwise and then the solution was warmed to room
temperature and stirred over 48 hours. The reaction was concentrated, diluted with ethyl acetate
(50 mL), washed with brine (50 mL) and the organic layer was then dried over sodium sulfate.
The product was filtered, concentrated in vacuo, and then recrystallized with ethyl acetate. The
product (0.214 g, 74% yield) was collected as a white solid.
1
H NMR (400 MHz, Chloroform-d)
δ
H
7.91 (s, 1H), 7.39-7.27 (m, 5H), 6.43 (t, J = 6.1 Hz, 1H), 4.44 (d, J = 6.0 Hz, 2H), 3.55 (d, J =
77.3 Hz, 2H), 2.87 (s, 3H).

N
H
N
H
O
N
OH
O
94

tert-butyl 1-benzylhydrazine-1-carboxylate (4.15). In a round bottom flask, with a stirbar and
an addition funnel affixed to the neck of the flask, benzylhydrazine hydrochloride (1.90 g, 12.00
mmol) and di-tert-butyl-dicarbonate (3.04 mL, 13.23 mmol) were stirred in THF (60 mL) at
room temperature for 15 minutes. Next, a solution of sodium bicarbonate (3.02 g, 35.9 mmol) in
40 mL of water was added dropwise over two hours via the addition funnel. The reaction was
allowed to stir at room temperature and monitored by TLC until the reaction was complete.
Once finished, the solution was concentrated in vacuo and then diluted with ethyl acetate (200
mL). The organic solution was washed with aqueous saturated sodium bicarbonate solution (1 x
200 mL), and then the aqueous layer was extracted with ethyl acetate (3 x 200 mL). The
combined organic layers were dried over sodium sulfate, filtered, and then concentrated in
vacuo. The crude product was purified via column chromatography run with a mobile phase of
30% ethyl acetate:hexane. The purified product was collected in 89% yield.
1
H NMR (400
MHz, Chloroform-d) δ
H
7.37-7.26 (m, 5H), 4.56 (s, 2H), 4.00 (s, 2H), 1.49 (s, 9H).

tert-butyl 1-benzyl-2-(benzylcarbamoyl)hydrazine-1-carboxylate (4.16). To a round bottom
flask was added a stirbar, the hydrazine starting material (4.15, 2.37 g, 10.66 mmol), and 15 mL
H
2
N
N O
O
N
H
N
H
O
N O
O
95
of DMF. The flask was cooled to 0
o
C and then 1,1’-carbonyldiimidazole (CDI) (1.90 g, 11.72
mmol) was added and the reaction was stirred for one hour at 0
o
C. Next, benzyl amine (1.16
mL, 10.6 mmol) and triethyl amine (1.64 mL, 11.76 mmol) were added to the solution and the
reaction was allowed to stir at room temperature overnight. The solvent was removed in vacuo,
and then the crude mixture was diluted with ethyl acetate (150 mL). The mixture was washed
with brine (150 mL), aqueous saturated sodium bicarbonate (150 mL), 1 M potassium bisulfate
solution (150 mL) and then brine again (150 mL). The organic layer was dried over magnesium
sulfate, filtered, and concentrated in vacuo. The white solid product was recrystallized with
ethyl acetate and hexane, producing the pure white crystalline product in 44% yield.
1
H NMR
(500 MHz, Chloroform-d) δ
H
7.32-7.25 (m, 9H), 7.18 -7.16 (m, 1H), 6.02 (s, 1H), 5.14 (t, J = 5.8
Hz, 1H), 4.56 (s, 2H), 4.32 (d, J = 6.1 Hz, 2H), 1.45 (s, 9H).

N,2-dibenzylhydrazinecarboxamide (4.17). To the protected hydrazine (4.16, 0.73 g, 2.05
mmol) in a round bottom flask was added a stirbar and 88 mL of DCM. The solution was stirred
at room temperature and then 4.5 mL (58.8 mmol) of trifluoroacetic acid was added to the flask.
The reaction was stirred at room temperature for approximately 5 hours, until the deprotection
showed completion via TLC. Next, the reaction mixture was concentrated in vacuo, diluted with
ethyl acetate (100 mL), washed with brine (100 mL), and dried over sodium sulfate. The organic
N
H
N
H
O
N
H
96
layer was filtered and then concentrated in vacuo. The product was used crude for the next
reaction without further purification or characterization.

N-benzyl-N-(3-benzylureido)glycine (4.8). The crude starting material (4.17, 0.816 g, 3.20
mmol) was diluted with 85 mL of methanol in a round bottom flask with a stirbar. Glyoxylic
acid monohydrate (0.882 g, 9.58 mmol) was added to the flask, and the reaction was stirred for
30 minutes at room temperature. Sodium cyanoborohydride (0.301 g, 4.79 mmol) was added
and the reaction was stirred for one hour at room temperature. Next, 2 N HCl solution was added
to the reaction mixture until a pH of 1 was reached. Saturated sodium bicarbonate solution was
added to adjust the pH to 4 and then the solution was extracted with DCM (3 x 100 mL). The
combined organic layers were washed with brine (300 mL) and then dried over sodium sulfate.
The organic layers were filtered and then concentrated in vacuo to give a crude white solid. The
crude product was recrystallized with ether to give pure white solid product in 45% yield.
1
H
NMR (500 MHz, DMSO-d
6
) δ
H
12.68 (s, 1H), 7.43-7.15 (m, 9H), 7.05 (d, J = 9.2 Hz, 1H), 6.84
(t, J = 6.4 Hz, 1H), 4.12 (d, J = 5.0 Hz, 2H), 3.89 (s, 2H), 3.50 (s, 2H).

N
H
N
H
O
N
OH O
97

2,2-diethoxy-N-(thiophen-2-ylmethyl)ethanamine (4.18). To a round bottom flask with a
stirbar was added aminoacetaldehyde diethyl acetal (0.55 mL, 3.78 mmol) and thiophene-2-
aldehyde (0.35 mL, 3.74 mmol). The reaction was stirred neat at 80
o
C for 20 minutes, before
cooling to room temperature and adding 3 mL ethanol and 0.142 g (3.75 mmol) of sodium
borohydride. The reaction was stirred at room temperature overnight and then concentrated in
vacuo. The crude product was diluted with ethyl acetate (50 mL), washed with water (50 mL),
and then dried over sodium sulfate before filtering and concentrating in vacuo. The crude
product was then purified by silica column chromatography eluting at 20% ethyl acetate: hexane
to yield 0.613 g (71%) of product as a colorless oil.
1
H NMR (500 MHz, Chloroform-d) δ
H
7.21
(ddd, J = 5.0, 1.3, 0.8 Hz, 1H), 6.94 (dddd, J = 7.8, 4.4, 3.4, 0.9 Hz, 2H), 4.61 (t, J = 5.6 Hz, 1H),
4.01 (s, 2H), 3.73-3.66 (m, 2H), 3.57-3.50 (m, 2H), 2.78 (d, J = 5.6 Hz, 2H), 1.21 (t, J = 6.8 Hz,
6H).
13
C NMR (126 MHz, Chloroform-d) δ 144.04, 126.59, 124.90, 124.37, 102.17, 62.40,
51.30, 48.36, 15.40.

HN
S
O
O
98

(9H-fluoren-9-yl)-methyl-(2-((2,2-diethoxyethyl)(thiophen-2-ylmethyl)amino)-2-
oxoethyl)carbamate (4.19). Fmoc-glycine (0.361 g, 1.21 mmol) was added to a vial alone with
DMF (2.87 mL) and a stirbar. Then, Hunig’s base (DIEA) (0.23 mL, 1.32 mmol) was added to
the solution, followed by HATU (0.508 g, 1.34 mmol). The reaction was stirred at room
temperature for 30 minutes, then a solution of the amine (4.18, 0.307 g, 1.34 mmol) in 1 mL of
DMF was added dropwise and the reaction was stirred overnight at room temperature. The
reaction was then concentrated in vacuo and purified via silica column chromatography eluted
with 50% ethyl acteate: hexane. The purified product (0.672 g, 99% yield) was collected as a
yellow oil.
1
H NMR (400 MHz, Chloroform-d) δ
H
7.76 (d, J = 7.7 Hz, 2H), 7.67-7.57 (m, 2H),
7.44-7.36 (m, 2H), 7.35-7.28 (m, 2H), 7.26-7.22 (m, 1H), 7.02-6.98 (m, 1H), 6.95 (dt, J = 5.1,
3.8 Hz, 1H), 5.89-5.69 (m, 1H), 4.68 (t, J = 5.3 Hz, 1H), 4.50 (t, J = 5.1 Hz, 1H), 4.38 (d, J = 7.3
Hz, 2H), 4.19 (t, J = 4.2 Hz, 2H), 3.72 (ddq, J = 21.1, 9.2, 7.0 Hz, 2H), 3.60-3.40 (m, 2H), 3.36
(d, J = 5.2 Hz, 1H), 1.21 (t, J = 9.1 Hz, 6H).

N
H
O
O
O
S
N
O
O
99

2-amino-N-(2,2-diethoxyethyl)-N-(thiophen-2-ylmethyl)acetamide (4.9). The protected
amine (4.19, 0.672 g, 1.32 mmol) was diluted with 10 mL DCM in a round bottom flask with a
stirbar. Next, diethyl amine (6 mL) was added at room temperature and the reaction was allowed
to stir overnight. Next, the mixture was concentrated in vacuo, diluted with ethyl acetate (50
mL), celite was added and concentrated again. The crude product deposited onto the celite was
dry loaded onto a silica column and eluted with a mobile phase of 50% ethyl acetate: hexane.
The purified product was collected in 53% yield.
1
H NMR (400 MHz, Chloroform-d) δ
H
7.18
(dd, J = 5.1, 1.2 Hz, 1H), 6.92-6.88 (m, 2H), 4.80 (s, 2H), 4.46 (t, J = 5.1 Hz, 1H), 4.06 (t, J =
6.7 Hz, 2H), 3.69-3.59 (m, 4H), 3.32 (d, J = 5.2 Hz, 2H), 1.00-0.85 (m, 6H). LC-MS: calculated
286.14, observed 287.1 [M+1]

(S)-(9H-fluoren-9-yl)-methyl-(1-((2,2-diethoxyethyl)(thiophen-2-ylmethyl)amino)-1-oxo-3-
phenylpropan-2-yl)carbamate (4.20). Compound 4.20 was synthesized and purified similarly
H
2
N
N
O
S
O
O
N
H
N O
S
O
O
O
O
100
to compound 4.19, in 86% yield.
1
H NMR (400 MHz, Chloroform-d) δ
H
7.76 (dt, J = 7.8, 1.0
Hz, 2H), 7.59-7.53 (m, 2H), 7.43-7.37 (m, 2H), 7.34-7.27 (m, 2H), 7.27-7.21 (m, 1H), 7.21-7.08
(m, 2H), 6.97-6.88 (m, 2H), 6.87-6.82 (m, 1H), 5.62 (d, J = 8.8 Hz, 1H), 5.04-4.89 (m, 1H), 4.58
(d, J = 15.0 Hz, 1H), 4.42-4.12 (m, 4H), 3.79-3.27 (m, 5H), 3.19 (dd, J = 15.5, 5.6 Hz, 1H), 3.11-
2.93 (m, 3H), 1.24-1.10 (m, 6H).

(S)-2-amino-N-(2,2-diethoxyethyl)-3-phenyl-N-(thiophen-2-ylmethyl)propanamide (4.10).
Compound 4.10 was synthesized and purified similarly to compound 4.9, in 89% yield.
1
H
NMR (400 MHz, Chloroform-d) δ
H
7.33-7.27 (m, 1H), 7.25-7.12 (m, 6H), 6.96-6.90 (m, 1H),
4.92 (d, J = 15.0 Hz, 1H), 4.68-4.55 (m, 2H), 4.34 (t, J = 5.3 Hz, 1H), 4.09-3.98 (m, 1H), 3.76-
3.65 (m, 1H), 3.65-3.52 (m, 2H), 3.43 (dqd, J = 9.1, 7.0, 2.0 Hz, 1H), 3.38-3.22 (m, 1H), 3.22-
3.08 (m, 2H), 3.02 (m, 1H), 2.84 (m, 1H), 1.17 (m, 6H). LC-MS: calculated 376.18, observed
377.2 [M+1].

H
2
N
N O
S
O
O
101

(S)-(9H-fluoren-9-yl)-methyl-(1-((2,2-diethoxyethyl)amino)-1-oxo-3-phenylpropan-2-
yl)carbamate (4.21). Compound 4.21 was synthesized and purified similarly to compound 4.19,
in 91% yield.
1
H NMR (500 MHz, Chloroform-d) δ
H
7.76 (d, J = 7.5 Hz, 2H), 7.57-7.51 (m,
2H), 7.40 (td, J = 7.5, 1.1 Hz, 2H), 7.35-7.27 (m, 4H), 7.27-7.16 (m, 3H), 5.88 (s, 1H), 5.43 (d, J
= 7.3 Hz, 1H), 4.43 (dd, J = 10.6, 7.1 Hz, 1H), 4.40-4.27 (m, 3H), 4.19 (t, J = 6.9 Hz, 1H), 3.60
(tq, J = 10.2, 7.1 Hz, 2H), 3.50-3.35 (m, 2H), 3.30 (d, J = 5.8 Hz, 2H), 3.15-2.98 (m, 2H), 1.14
(td, J = 7.0, 4.7 Hz, 6H).
13
C NMR (126 MHz, Chloroform-d) δ 170.83, 143.85, 141.43, 129.43,
128.84, 127.89, 127.22, 125.22, 125.13, 120.14, 100.62, 67.21, 63.06, 56.54, 47.27, 42.00, 39.03,
15.37.

(S)-2-amino-N-(2,2-diethoxyethyl)-3-phenylpropanamide (4.11). Compound 4.11 was
synthesized and purified similarly to compound 4.9, in 27% yield.
1
H NMR (400 MHz,
Chloroform-d) δ
H
7.40 (q, J = 4.7, 2.8 Hz, 1H), 7.32-7.28 (m, 2H), 7.26-7.19 (m, 3H), 4.46 (t, J
= 5.4 Hz, 1H), 3.68 (dqd, J = 9.4, 7.1, 1.4 Hz, 2H), 3.61 (dd, J = 9.3, 4.2 Hz, 1H), 3.51 (dqd, J =
H
N
N
H
O
O
O
O
O
H
2
N
N
H
O
O
O
102
9.5, 7.0, 4.8 Hz, 2H), 3.39 (t, J = 5.7 Hz, 2H), 3.25 (dd, J = 13.7, 4.2 Hz, 1H), 2.69 (dd, J = 13.7,
9.3 Hz, 1H), 1.20 (tdd, J = 7.1, 2.1, 0.6 Hz, 6H).

(S)-N,2-dibenzyl-2-(2-((1-((2,2-diethoxyethyl)amino)-1-oxo-3-phenylpropan-2-yl)amino)-2-
oxoethyl)hydrazinecarboxamide (4.22). Compound 4.22 was synthesized similarly to
compound 4.19, and used crude for the next reaction without further purification or
characterization.

N
H
N
H
O
N
HN
O
NH O
O
O
103

N,2-dibenzyl-2-(2-((2-((2,2-diethoxyethyl)(thiophen-2-ylmethyl)amino)-2-oxoethyl)amino)-
2-oxoethyl)hydrazinecarboxamide (4.23). Compound 4.23 was synthesized similarly to
compound 4.19, and used crude for the next reaction without further purification or
characterization.

(S)-N-benzyl-2-(2-((1-((2,2-diethoxyethyl)(thiophen-2-ylmethyl)amino)-1-oxo-3-
phenylpropan-2-yl)amino)-2-oxoethyl)-2-methylhydrazinecarboxamide (4.24). The
carboxylic acid (4.7, 0.091 g, 0.38 mmol) was dissolved in 2 mL of DMF in a vial with a stirbar.
Next, Hunig’s base (DIEA) (0.074 mL, 0.42 mmol) was added followed by HATU (0.16 g, 0.42
mmol). The reaction was stirred at room temperature for 30 minutes and then the amine (4.10,
0.159 g, 0.42 mmol) was added to the reaction mixture. The reaction was stirred at room
N
H
N
H
O
N
HN
O
N O
S
O
O
N
H
N
H
O
N
N
H
O
N O
S
O
O
104
temperature overnight before concentrating in vacuo and diluting with ethyl acetate (50 mL).
The solution was washed with brine (2 x 50 mL) before drying the combined organic layers over
sodium sulfate. The solution was filtered, concentrated in vacuo, and used crude for the next
step without further purification or characterization.

(6S)-N,2,6-tribenzyl-4,7-dioxooctahydro-1H-pyrazino[2,1-c][1,2,4]triazine-1-carboxamide
(4.1). Compound 4.1 was synthesized and purified similarly to compound 4.3, in 8% yield. The
product was characterized via LC-MS. LC-MS: calculated 483.23, observed 482.2 [M-1].

N,2-dibenzyl-4,7-dioxo-8-(thiophen-2-ylmethyl)octahydro-1H-pyrazino[2,1-
c][1,2,4]triazine-1-carboxamide (4.2). Compound 4.2 was synthesized and purified similarly to
N
N
N
NH
O NH
O
O
N
N
N
N
O NH
O
O
S
105
compound 4.3, in 7% yield.
1
H NMR (600 MHz, Acetone-d
6
) δ
H
7.49 (t, J = 5.6 Hz, 1H), 7.41
(dd, J = 5.1, 1.2 Hz, 1H), 7.34 (dd, J = 6.7, 3.1 Hz, 2H), 7.30-7.28 (m, 5H), 7.24 (ddd, J = 8.3,
4.7, 1.8 Hz, 3H), 7.12 (dd, J = 3.5, 1.2 Hz, 1H), 7.01 (dd, J = 5.1, 3.4 Hz, 1H), 5.82 (dd, J = 10.4,
4.1 Hz, 1H), 4.92-4.74 (m, 2H), 4.69 (d, J = 18.3 Hz, 1H), 4.51-4.23 (m, 3H), 4.22 (s, 2H), 3.92-
3.84 (m, 2H), 3.51 (dd, J = 11.7, 4.2 Hz, 1H), 3.49-3.44 (m, 1H).
13
C NMR (100 MHz, Acetone-
d
6
) δ 164.59, 164.44, 156.13, 154.11, 152.23, 141.71, 141.10, 139.97, 137.30, 130.22, 129.36,
129.12, 128.63, 128.14, 128.02, 127.56, 126.77, 123.89, 79.60, 64.16, 61.54, 54.11, 51.10, 46.15,
44.84, 44.28. LC-MS: calculated 489.18, observed 488.2 [M-1].

(6S)-N,6-dibenzyl-2-methyl-4,7-dioxo-8-(thiophen-2-ylmethyl)octahydro-1H-pyrazino[2,1-
c][1,2,4]triazine-1-carboxamide (4.3). To a round bottom flask equipped with a stirbar was
added the crude starting material (4.24, 0.233 g, 0.39 mmol) and 5 mL of formic acid. The
reaction was stirred at room temperature overnight. The reaction was then cooled to 0
o
C and
diluted with acetone, and the pH was adjust to 4 using 2N NaOH. The acetone was then
removed in vacuo, and the product was extracted with ethyl acetate (3 x 50 mL). The combined
organic layers were dried over sodium sulfate, filtered, and then concentrated onto celite and
purified via silica column chromatography. The purified product was eluted with a pure ethyl
N
N
N
N
O NH
O
O
S
106
acetate, followed by 10% methanol: ethyl acetate. The purified product (0.02 g, 0.04 mmol) was
collected as a pale yellow oil in 10% yield.
1
H NMR (600 MHz, Acetone-d
6
) δ
H
7.45 (t, J = 6.4
Hz, 1H), 7.35 (dd, J = 5.1, 1.2 Hz, 1H), 7.30 (t, J = 7.6 Hz, 2H), 7.27-7.24 (m, 2H), 7.21 (t, J =
7.2 Hz, 1H), 7.14-7.08 (m, 5H), 7.02-6.99 (m, 1H), 6.93 (dd, J = 5.1, 3.5 Hz, 1H), 5.25 (dd, J =
10.8, 4.1 Hz, 1H), 5.14 (t, J = 5.6 Hz, 1H), 4.94 (d, J = 15.0 Hz, 1H), 4.54 (d, J = 15.0 Hz, 1H),
4.30 (ddd, J = 73.4, 15.2, 6.3 Hz, 2H), 3.76 (t, J = 11.2 Hz, 1H), 3.49 (d, J = 16.9 Hz, 1H), 3.36-
3.24 (m, 3H), 3.11 (d, J = 16.9 Hz, 1H), 2.76 (s, 3H).
13
C NMR (151 MHz, Acetone-d
6
) δ
166.67, 163.32, 156.52, 141.43, 139.92, 137.62, 130.42, 129.21, 129.12, 128.03, 127.89, 127.69,
127.60, 127.43, 126.69, 61.38, 57.03, 55.93, 50.99, 45.59, 45.33, 44.17, 36.95. LC-MS:
calculated 503.20, observed 502.1 [M-1].

107
4.5 Chapter 4 References

1
Xu, C.; Bailly-Maitre, Reed, J. C. J. Clin. Invest. 2005, 115, 2656.

2
Ni, M.; Lee, A. S. FEBS Lett. 2007, 581, 3641.

3
Pfaffenbach, K. T.; Lee, A. S. Curr. Opin. Cell Biol. 2011, 23, 150.

4
Luo, S.; Mao, C.; Lee, B.; Lee, A. S. Mol. Cell. Biol. 2006, 26, 5688.

5
Oyadomari, S.; Mori, M. Cell Death Differ. 2004, 11, 381.

6
Li, J.; Lee, A. S. Curr. Mol. Med. 2006, 6, 45.

7
Rutkowski, D. T.; Kaufman, R. J. TRENDS Cell Biol 2004, 14, 20.

8
Reddy, R. K.; Mao, C.; Baumeister, P.; Austin, R. C.; Kaufman, R. J.; Lee, A. S. J. Biol. Chem.
2003, 278, 20915.

9
Hajnokzky, G.; Davies, E.; Madesh, M. Biochem. Biophys. Res. Comm. 2003, 304, 445.

10
Hammadi, M.; Oulidi, A.; Gackiere, F.; Katsogiannou, M.; Slomianny, C.; Roudbaraki, M.;
Dewailly, E.; Delcourt, P.; Lepage, G.; Lotteau, S.; Ducreux, S.; Prevarskaya, N.; Van
Coppenolle, F. FASEB J. 2013, 27, 1600.

11
Lee, S. A. Methods 2005, 35, 373.

12
Pyrko, P.; Schonthal, A. H.; Hofman, F. M.; Chen, T. C.; Lee, A. S. Cancer Res. 2007, 67,
9809.

13
Fernandez, P. M.; Tabbara, S. O.; Jacobs, L. K.; Manning, F. C. R.; Tsangaris, T. N.;
Schwartz, A. M.; Kennedy, K. A.; Patierno, S. R. Breast Cancer Res. Treat. 2000, 59, 15.

14
Daneshmand, S.; Quek, M. L.; Lin, E.; Lee, C.; Cote, R. J.; Hawes, D.; Cai, J.; Groshen, S.;
Lieskovsky, G.; Skinner, D. G.; Lee, A. S.; Pinski, J. Human Pathol. 2007, 38, 1547.

15
Lee, E.; Nichols, P.; Spicer, D.; Groshen, S.; Yu, M. C.; Lee, A. S. Cancer Res. 2006, 66,
7849.

16
Lee, A. S. Cancer Res. 2007, 67, 3496.

17
Arap, M. A.; Lahdenranta, J.; Mintz, P. J.; Hajitou, A.; Sarkis, A. S.; Arap, W.; Pasqualini, R.
Cancer Cell 2004, 6, 275.

108

18
Wang, M.; Wey, S.; Zhang, Y.; Ye, R.; Lee, A. S. Antioxid. Redox Signal. 2009, 11, 2307.

19
Misra, U. K.; Deedwania, R.; Pizzo, S. V. J. Biol. Chem. 2006, 281, 13694.

20
Shani, G.; Fischer, W. H.; Justice, N. J.; Kelber, J. A.; Vale, W.; Gray, P. C. Mol. Cell Biol.
2008, 28, 666.

21
Burkhanov, R.; Zhao, Y.; Goswami, A.; Qui, S.; Schwarze, S. R.; Rangnekar, V. M. Cell
2009, 138, 377.

22
Davidson, D. J.; Haskell, C.; Majest, S.; Kherzai, A.; Egan, D. A.; Walter, K. A.; Schneider,
A.; Gubbins, E. F.; Solomon, L.; Chen, Z.; Lesniewsky, R.; Henkin, J. Cancer Res. 2005, 65,
4663.

23
Misra, U. K.; Gonzalez-Gronow, M.; Gawdi, G.; Wang, F.; Pizzo, S. V. Cell. Signal. 2004, 16,
929.

24
Goldenberg-Cohen, N.; Raiter, A.; Gaydar, V.; Dratviman-Storobinsky, O.; Goldstein, T.;
Weizman, A.; Hardy, B. Apoptosis 2012, 17, 278.

25
Paton, A. W.; Beddoe, T.; Thorpe, C. M.; Whisstock, J. C.; Wilce, M. C. J.; Rossjohn, J.;
Talbot, U. M.; Paton, J. C. Nature 2006, 443, 548.

26
Belvisi, L.; Colombo, L.; Manzoni, L.; Potenza, D.; Scolastico, C. Synlett 2004, 9, 1449.

27
McMillian, M.; Kahn, M. Drug Disc. Today 2005, 10, 1467.

28
Piergentili, A.; Del Bello, F.; Gentili, F.; Giannella, M.; Quaglia, W.; Vesprini, C.; Thomas, R.
J.; Robertson, G. M. Tetrahedron 2007, 63, 12912.

29
Oh, S.-W.; Lee, J. B.; Kim, B.; Jeon, S.; Kim, M.-K.; Nam, K.-H.; Ha, J.-R.; Bhatia, M.; Oh,
G. T.; Kim, D.-Y. Arch. Pharm. Res. 2012, 35, 1979.

109
Conclusions
A novel ω-alkynyl-docosahexaenoic acid was successfully synthesized for the use as a
clickable imaging probe and for further biological studies. Prelimanary enzymatic assays using
15-lipoxygenase were successfully run to confirm that the metabolism of the potential imaging
probe would be unaffected by the terminal alkyne tag.

Building blocks towards the synthesis of lipoxin analogs were synthesized successfully.
Chiral pools, such as L-rhamnose and S-glycidol, provided stereocenters for the synthesis of
target pieces.

Reverse-turn peptidomimetics were successfully designed and synthesized for the
potential modulation of the ER stress master regulator protein GRP78. The novel
hexahydropyrazinotriazinediones were designed for the investigation of structure-activity-
relationships between the aromatic ring structure placement on the compounds and biological
activity.
110
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115
Appendix: Selected Spectra
2.3

TMS
OH

!
" # $ $ $
$
$
$ $

116
2.2

TMS
Br

!
" # $ % !
# &
' ( !

) *

+ " ,

117
2.4

COOMe
TMS

!
" # " $
% & ' $
(

) ( %
* $

+
% ,
) "
#
-
%

!
"
# $
% & ' $ !
(

) ( %
* $

118
2.6

COOMe
TMS

! "
#

"
$ % & ' ( )

) *
+ )
( ( + )
,
)
- ) .
- )

) /
)
) ) )
) ) ) )
)

119
2.7

COOMe

120
2.9

COOMe

121
2.1

COO
-
Na
+

122
2.1

COO
-
Na
+

123
2.11

COO
-
OH
124
3.7

125
3.8

126
3.9

127
3.1

128
3.10

129
3.2

130
3.11

131
3.12

132
3.13

133
3.14

! " # $ $
$
$ $ $

$

$
$
$
$
134
3.3

135
3.15

136
3.16

137
3.17

138
3.18

139
3.4

140
3.20a

O
O O O
O
TMS

141
3.20b

O
O O O
O
TMS

! "

142
3.22

!

143
3.23

144
3.24

145
3.25

!

146
4.12

N
H
N
H
O
N O
O

147
4.13

N
H
N
H
O
NH

148
4.14

N
H
N
H
O
N
O
O

149
4.7

N
H
N
H
O
N
OH
O

150

4.15

H
2
N
N O
O

151
4.16

N
H
N
H
O
N O
O

152
4.8

N
H
N
H
O
N
OH O

153
4.18

HN
S
O
O

154
4.19

N
H
O
O
O
S
N
O
O

! "
#
$
%

155
4.10

H
2
N
N O
S
O
O

!
"
#
$

156
4.21

H
N
N
H
O
O
O
O
O

!
" #
$
%

157
4.11

H
2
N
N
H
O
O
O

158
4.1

N
N
N
NH
O NH
O
O
MS Spectrum
m/ z 100 200 300 400 500 600
0
20
40
60
80
100
*MSD2 SPC, time=6.102 of C:\CHEM32\1\DATA\ANNEMARIE\AMF1-283-4.D ES-API, Neg, Scan, Frag: 5
Max: 10568
601.0
429.9
484.1
546.1
113.0
483.3
600.0
487.3 486.2
529.3 528.1
482.2
Print of window 80: MS Spectrum
Instrument 1 10/24/2013 1:39:05 PM Marcos Sainz Page 1 of 1
159
4.2

N
N
N
N
O NH
O
O
S

!
"
#
$

160
4.2

N
N
N
N
O NH
O
O
S

161
4.2

N
N
N
N
O NH
O
O
S
MS Spectrum
m/ z 100 200 300 400 500
0
20
40
60
80
100
*MSD2 SPC, time=6.549 of C:\CHEM32\1\DATA\ANNEMARIE\AMF1-279-PTLC-2.D ES-API, Neg, Scan, Frag: 5
Max: 8114
249.0
524.1
550.9
446.3
490.0
536.2
507.1
445.3
489.2
535.1
113.0
488.2
444.2
534.1
Print of window 80: MS Spectrum
Instrument 1 10/24/2013 1:40:53 PM Marcos Sainz Page 1 of 1
162

4.3

N
N
N
N
O NH
O
O
S

!
"
#
$
%
&
'

163
4.3

N
N
N
N
O NH
O
O
S

164
4.3

N
N
N
N
O NH
O
O
S
MS Spectrum
m/ z 100 200 300 400 500
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Instrument 1 10/24/2013 1:41:55 PM Marcos Sainz Page 1 of 1

Abstract (if available)

Abstract The research presented within this dissertation is a compilation of three projects with the underlying theme of designing and synthesizing relevant molecules for biological studies. ❧ The first chapter serves as an introduction to lipid mediators and their role in inflammation. A brief overview of the lipid mediator pathways and their significance are discussed within chapter one. ❧ Chapter two introduces click chemistry as a tool for probing lipid-related biological pathways. The design and total synthesis of an alkynyl-docosahexaenoic acid analog for potential use as an imagining probe using click chemistry is described. Preliminary enzymatic assays were also performed to assess the stability and metabolism of the final product. ❧ The third chapter presents the design and synthesis of building blocks for the total synthesis of lipoxin analogs. The building block design was focused on incorporating moieties that would potentially be metabolically stable and biologically active upon later completion of the final structure. ❧ Chapter four introduces the protein GRP78 and its significance to cancer and disease development. The concept of reverse-turn peptidomimetics is briefly introduced and applied to the design of our target molecules. Novel potential modulators of GRP78 are successfully synthesized for biological testing.

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