Design & Synthesis of Prostaglandin Ethanolamide Covalent Probes for the Investigation of Novel Physiological Activity and Discovery of a Prostaglandin Intermediate Scaffold for FAAH Inhibition

A dissertation presented

by

Erin Laine Shelnut

to

The Department of Chemistry and Chemical Biology

In partial fulfillment of the requirements for the degree of Doctor of Philosophy

in the field of

Chemistry

Northeastern University Boston, Massachusetts June, 2012

Design and Synthesis of Prostaglandin Ethanolamide Covalent Probes for the Investigation of Novel Physiological Activity and Discovery of a Prostaglandin Intermediate Scaffold for FAAH Inhibition

by

Erin L. Shelnut

ABSTRACT OF DISSERTATION

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the field of Chemistry in the Graduate School of Northeastern University June, 2012

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Abstract

Prostaglandin ethanolamides (Prostamides) are an emerging class of endogenous

eicosanoids derived from cyclooxygenase (COX) metabolism of the endocannabinoid,

anandamide. The chemical structure of the prostamides resembles that of the

prostaglandins, with the main distinction attributed to an ethanolamide group in place of the carboxylic acid group. Their biological function has yet to be resolved, however the biosynthetic route for prostamide formation has been well established and their existence in vivo postulated.

While the structures and biosynthetic routes are indeed similar, there is growing evidence that prostamides and their prostaglandin counterparts exhibit diverse biological actions. The substitution of ethanolamide for carboxylic acid is sufficient to give the prostamides a diverse biological profile. Indeed, the biosynthetic precursors of prostamides and prostaglandins share this chemical distinction, and are well-known to incite distinct processes. Anandamide is capable of activating the CB1 cannabinoid and TRPV1 vanilloid receptors and serves as their main endogenous substrate. Arachidonic acid, on the other hand, is inactive at both of these receptors and functions mainly as a secondary messenger in cellular signaling and as a precursor to the eicosanoid compounds. In fact, the major pathway of anandamide cannabinergic inactivation is hydrolytic catalysis by the enzyme fatty acid amide hydrolase (FAAH) to arachidonic acid. The disparity between the functions of arachidonic acid and anandamide give credibility to the hypothesis that prostamides might also possess a unique biological purpose.

Prostaglandins are known to act at well characterized prostaglandin receptors, named for the specific prostaglandin they bind. Prostamides similarly may interact with distinct, yet to be characterized receptors. Our laboratory attempts to address this prospect by development of novel prostamide analogs for screening against known and orphan receptors and for in vivo inflammation and immunomodulation studies.

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Chapter 1 provides an overview of the prostamide biosynthetic pathway, including

detailed discussion of anandamide release and action and cyclooxygenase structure and

function. Additionally, a comprehensive review of current prostamide research covering

stability and pharmacology in support of, and in opposition to, the functional distinction

between the prostamides and prostaglandins is presented. Conflicting opinions exist as to the uniqueness of the prostamides’ pharmacology, and exploring both sides of the dispute gives a comprehensive review of studies attempting to elucidate prostamides’ physiological targets. Finally, their emerging role in inflammation, neuroinflammation and neuroplasticity is addressed. Most strikingly, evidence suggests that neuroprotective qualities long attributed to the actions of the endocannabinoid, anandamide, are instead the result of its oxidized metabolite, prostamide.

Chapter 2 presents the design and synthesis of a set of prostamide analogs to be screened against existing known and orphan receptors and to undergo inflammation and immunomodulation studies. Analogs are designed to incorporate tail moieties that covalently bind a target receptor protein. Modification in head group functionality attempt to address issues of prostamide metabolic stability, and variation of the stereochemistry of the 15R hydroxyl may lead to optimization of a possible hydrogen bond with the active site. The challenges addressed in this extensive synthesis include instability of the cyclopentyl ring hydroxy groups and unachievable chiral reduction of the 15-keto group. The assignment of resulting chirality is designated by thorough Mosher analysis. Using the synthetic route established here, the tail hydroxyl moiety can serve as an entrance to a vast number of future alterations in functionality of both head and tail substituents.

The results of biological screening of prostamide analogs on well-established and orphan receptors are presented in Chapter 3. None of the isolated receptors screened showed significant activity toward the prostamides tested. Additionally described within this chapter is the effect of prostamide treatment on an in vivo model of murine inflammatory

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peritonitis. The single analog tested showed enhanced clearance of pro-inflammatory

polymorphonuclear neutrophils and thus exhibited behavior consistent with resolution of

inflammation. This study also revealed the inability of the prostaglandin probe to activate

the EP prostaglandin receptors. Finally, prostamide’s immunosuppressive effects similar to

those of anandamide were demonstrated.

Chapter 4 outlines areas of study in which prostamide pharmacological activity

can be further explored. Investigations of prostamide action in cells of the central nervous

system represent the most promising course of research. Prostamide E2 enhances

neuroprotection and suppresses cytokine production, and due to increased production of the precursor anandamide, is expected to be present at relevant concentration in the brain. A study of prostamide activity at EP receptor splice variants may be integral in determining prostamide’s unique pharmacology. Screening prostamide against nuclear receptors such as

PPARγ has yet to be completed. Due to the ability of PPARγ to bind prostaglandins and fatty

acids, screening on this receptor will determine whether prostamide is an endogenous

ligand. Isolation of prostamide-selective receptors from tissues sensitive to prostamide

treatment can be achieved by employing radiolabelled or biotinylated, bifunctional

prostamide probes. Once isolated, the prostamide-bound proteins can undergo mass

spectrometry studies to aid in characterization. Finally, design and synthesis of a

subsequent set of prostamide analogs is proposed within this chapter.

Chapter 5 introduces a secondary project in which three intermediates of

prostamide synthesis were discovered to exhibit inhibition of the FAAH enzyme. The enone

and the S and R allylic alcohol compounds inhibited of FAAH in the range of 100 nM-1 µM.

Thus, this set of bicyclic lactones represents a promising foundation from which to optimize

FAAH inhibition employing SAR study. An initial set of compounds including modifications at

the 11-hydroxyl group, at the 20-hydroxyl group, and of the chain length between the 15-

hydroxyl and ω tail were designed to assess the structure-activity relationship of the

-5- inhibitors to rFAAH. The foundations of a pharmacophore for these bicyclic lactone FAAH inhibitors were roughly established. The 15S-hydroxyl, 20-(1-adamantane carboxylate) compound AM7666, gave the greatest inhibition of the designed analogs, and its large bulk and hydrophobicity suggests that the binding pocket for the lipophilic portion of the inhibitors is considerably large.

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Acknowledgements

Through the enormous challenge of completing this dissertation I realize I owe enormous

gratitude to several members of the Department of Chemistry & Chemical Biology and the

Center for Drug Discovery. Thank you for providing me this opportunity to learn invaluable research skills and grow as an independent researcher.

I am exceeding thankful to Dr Alexandros Makriyannis for his guidance and investment.

Thank you for recognizing great potential in me and presenting me your most challenging and interesting research project. Throughout the years you have instilled in me a resilience

I will carry always, and I am very grateful for it.

I attribute my thanks to all of the members of the Center for Drug Discovery, both past and present, including Dr Lakshmipathi Pandrinathan for his invaluable guidance and training in proper methods of organic synthesis, Dr Kumara Vadivel Subramanian for his continuous assistance in all things lab related, and Jodi Wood for her patience and understanding in training me to perform biological assays. Thank you for sharing your knowledge and investing your time to aid me in my research pursuits and grant me the skills to become a better scientist.

I am infinitely grateful to the members of the CDD office, Shawntelle Dillon, Sarah

Strassburger, and Brett Greene. Even on your busiest days, you made time to help me with various requests and even gave me emotional support when I needed it most.

A number of collaborators have contributed to the research objectives described in this dissertation. Dr Bryan Roth at the University of North Carolina – Chapel Hill screened our compounds on known and orphan receptors through a Psychoactive Drug Screening

Program funded by the National Institute of Mental Health. Dr Charles Serhan and members

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of his group at Harvard Medical School, Drs Rong Yang and Nan Chiang, designed and performed the mouse peritonitis inflammatory studies and the prostaglandin EP receptor

ECIS studies. Dr Toby Eisenstein and Joseph Meissler at Temple University undertook the testing of our compounds on modulation of the immune system. These contributions were integral to the completion of this dissertation.

I am appreciative to the members of my dissertation committee, Drs Robert Hanson, Sunny

Zhou, and George O’Doherty for investing their time and assistance in the completion of this dissertation.

Support for this research was provided by a training grant from the National Institute of

Health.

Finally, to my Mom and Dad, thank you for your unwavering love and support. I could not have dreamed of making it this far without you having my back. You supplied endless encouragement and helped me to persevere in my most difficult times. To Patrick, thank you for surviving this with me. You have been my rock, a cool and calming force when I needed. This accomplishment would not have been possible without the three of you in my life. Thank you.

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Table of Contents

Abstract 2

Acknowledgements 7

Table of Contents 9

List of Abbreviations 15

1 Biosynthesis and Pharmacology of Prostaglandin 21 Ethanolamides (Prostamides)

Part A: Prostamide Biosynthesis 23

1.1 Biosynthesis, Action, and Degradation of the Prostamide 23 Precursor, Anandamide

1.2 Oxidative Metabolism of Fatty Acids and Fatty Acid Amides by 26 Cyclooxygenase Enzymes

1.2.1 Mechanism of Fatty Acid Oxidation 28

1.2.1.1 Autoxidation of Fatty Acids 28

1.2.1.2 Cyclooxygenase Crystal Structure and Active Site Interactions 29

1.2.1.3 Catalysis of Fatty Acid Oxidation by Cyclooxygenase 31

1.2.2 Differentiation of the Cyclooxygenase-1 and Cyclooxygenase-2 33 Isoforms

1.2.3 Inhibition of Cyclooxygenase 36

Part B: Prostamide Pharmacology 39

1.3 Debate Concerning Hydrolysis of Prostamides to Prostaglandins 42

1.3.1 Evidence of Negligible Prostamide Hydrolysis 42

1.3.2 Evidence of Substantial Prostamide Hydrolysis 46

1.4 Debate Concerning Distinction between the Biological Actions of 51 Prostamides and Prostaglandins

1.4.1 Evidence of Prostamide Biological Actions Distinct from those of 51 Prostaglandin

1.4.1.1 Isolated Receptor and Functional Tissue Studies 52

1.4.1.2 Confocal Microscopy Studies 57

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1.4.1.3 Prostamide Selective Antagonists 60

1.4.1.4 Cells that Express Prostamide-Selective Receptors Alone 65

1.4.1.5 Differences in Ophthalmological Pharmacology 66

1.4.2 Evidence of Prostamide Biological Actions Identical to Those 66 of Prostaglandin

1.4.2.1 Isolated Receptor and Functional Tissue Studies 67

1.4.2.2 Super-Sensitivity and Super-Coupling of FP Receptors in Tissue 68

1.4.2.3 FP Receptor Knockout 68

1.4.2.4 Confocal Microscopy Studies 69

1.4.2.5 Similarities in Ophthalmological Pharmacology 70

1.4.3 The Compromise: FP Receptor Splice Variants 71

1.5 Prostamide Action on Established Proteins 76

1.5.1 Receptors that Recognize Anandamide 76

1.5.2 Peroxisome Proliferation-Activated Receptor 78

1.5.3 Lipid Metabolizing Enzymes 80

1.6 Prostamides in Inflammation and Other Systems 83

1.6.1 Presence of Prostamide at Physiologically Relevant Concentration 83 In Vivo

1.6.2 Prostamides in Inflammation, Neuroinflammation and 84 Neuroplasticity

1.6.3 Misidentification of Prostamides as Prostaglandins 88

References 91

2 Design and Synthesis of α-Head and ω-Tail Modified 101 Prostamide E2 Analogs

2.1 Analog Design and Rationale 102

2.1.1 Configuration about the Cyclopentyl Ring 103

2.1.2 Covalent ω-Tail Modifications as Affinity Probes 103

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2.1.2.1 The Photoactivatable Azido and the Electrophilic Isothiocyanato 103 Groups

2.1.2.2 Utility of Covalent Affinity Probes 105

2.1.3 α-Head Group Modifications 107

2.1.4 The 15-Hydroxyl Stereocenter 108

2.2 Prostamide Analog Synthesis 111

2.2.1 Challenges in Prostaglandin and Prostamide Synthesis 111

2.2.2 Novelty and Advantage of the Current Synthesis 113

2.2.3 Synthetic Steps and Related Mechanisms 118

2.2.3.1 Synthesis of the ω-Chain 118

2.2.3.1.1 Chiral Reduction of the 15-Keto Group 120

2.2.3.1.2 Mosher Analysis of the 15-Hydroxyl Stereochemistry 121

2.2.3.1.3 Attempts at Chiral Reduction of the 15-Keto Group 127

2.2.3.2 Synthesis of the α-Chain 131

2.2.3.3 Functionalization of the ω-Tail 133

2.2.3.4 Functionalization of the α-Head Amide 135

2.2.3.5 Synthesis of the Non-Functionalized ω-Chain and Glycerol Ester 139 Analogs

References 141

3 Evaluation of Prostamides by Receptor Screening and as 146 Lipid Mediators in Inflammation and Immunomodulation

3.1 Biological Screening of Prostamides on Established and 147 Orphan Receptors

3.2 Effects of Prostamide on Inflammation 153

3.2.1 Reduction of Polymorphonuclear Neutrophils by Prostamide 154 Treatment of Zymosan A-Induced Inflammation

3.2.2 Determination of G-protein signaling pathways by Electric Cell- 156 Substrate Impedance Sensing

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3.2.3 Implementation of ECIS to Determine the Activity of Prostamide 161 on the Prostaglandin Receptors

3.3 Effects of Prostamide on Immunosuppression 163

3.4 Summary of Results 168

References 170

4 Future Directions for Prostamide Research 172

4.1 General Methods for Isolating Prostamide-Selective Target Proteins 174 from Prostamide-Sensitive Tissue

4.1.1 Binding Specificity Assay 174

4.1.2 Bifunctional Prostamide Probes 176

4.1.2.1 Isolation of Prostamide-Selective Proteins using Radiolabeled Probes 176

4.1.2.2 Emission Topography Imaging using Radiolabeled Probes 179

4.1.2.3 Affinity Purification of Prostamide-Selective Proteins using 180 Biotinylated Probes

4.2 Prostamide E2 Activity in the Central Nervous System 181

4.2.1 Neuroprotection by Prostamide E2 182

4.2.2 Interleukin-12 Suppression by Prostamide E2 183

4.2.3 Interleukin-2 Suppression by Prostamide E2 184

4.3 Prostamide Activity in Functional Tissue Assays 185

4.4 EP Receptor Splice Variants as the Putative Prostamide Receptor 186

4.5 Further Screening of Prostamide Analogs 188

4.5.1 Screening on PPARγ and Other Nuclear Receptors 188

4.5.2 Screening on Primary Cell Systems 189

4.6 Prostamide Activity in the Resolution of Inflammation 190

4.7 Suppression of the Immune System by Prostamide versus 191 Anandamide

4.8 Synthesis of Additional Prostamide Analogs for Biological Testing 192

4.8.1 Design and Proposed Synthesis of ω-Tail Modifications to Enhance 193 Binding Affinity

4.8.2 α-Head Modifications to Increase Metabolic Stability and 198 Incorporate Radio- and Biotin Labels

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4.8.2.1 Design and Proposed Synthesis of Radiolabeled Prostamide 198 Analogs

4.8.2.2 Design and Proposed Synthesis of Biotinylated Prostamide 200 Analogs

References 205

5 Inhibition of Fatty Acid Amide Hydrolase by a Novel Class 208 of Prostamide Intermediates

5.1 FAAH Enzyme Structure and Function 209

5.2 Inhibition of FAAH 212

5.2.1 Substrate-Based, Nonselective Inhibitors 212

5.2.2 Selective, Reversible Inhibitors 213

5.2.3 Selective, Irreversible Inhibitors 215

5.3 Rat FAAH vs Human FAAH: Differences in Inhibitor Sensitivity 218 Profiles

5.4 Discovery of Lead FAAH Inhibitors from Prostamide Intermediates 221

5.5 Design and Synthesis of Analogs of the Prostamide Intermediates 225 for FAAH Inhibition

5.6 Evaluation of the Prostamide Intermediate Analogs as FAAH 229 Inhibitors

5.7 Future Directions for FAAH Inhibition by Prostamide Intermediates 232

5.7.1 Molecular Modeling for Clarification of the Binding Site Interactions 232

5.7.2 Biochemical Assays to Assess Reversibility, Competitiveness, and 233 Selectivity

5.7.3 Synthetic Routes for an Extended Series of Prostamide 236 Intermediate Test Compounds

References 239

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Supplemental Information

Appendix 1: Supplemental Synthetic Methods and Characterization 246 for Compounds Presented in Chapter 2

Appendix 2: Biological Experimental Procedures and Supplemental 294 Data for Assays Presented in Chapter 3

Appendix 3: Supplemental Synthetic Methods and Characterization 304 for Compounds Presented in Chapter 4

Appendix 4: Biological Experimental Procedures for Assays Presented 327 in Chapter 4

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List of Abbreviations

2AG 2-Arachidonoyl Glycerol

AA Arachidonic Acid

ABHD4 α/β-Hydrolase-4

ACh Acetylcholine

AEA N-Arachidonoylethanolamide, Anandamide

AIBN Azobisisobutyronitrile altFP Splice Variant of Prostaglandin F receptor (FP) altFP4 Splice Variant of Prostaglandin F receptor (FP) 4

B-cells Bursa-derived cells

BFA Bimatoprost Free Acid

BINOL 1,1’-Bi-2-naphthol cAMP Cyclic Adenosine Monophosphate

Ca-TA Calcium-dependent Transacylase

CB Cannabinoid Receptor

CB1 Cannabinoid Receptor 1

CB2 Cannabinoid Receptor 2

CBS Corey-Bakshi-Shibata Chiral Catalyst

CBz Carboxybenzyl

CDI Carbonyldiimidazole

CHO Chinese Hampster Ovary Cells

CNS Central Nervous System

COX Cyclooxygenase

COX-1 Cyclooxygenase 1

COX-2 Cyclooxygenase 2

CREAE Chronic Relapsing-Remitting Experimental Autoimmune Encephalomyelitis

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Cyr61 Cystein-Rich Angiogenic Inducer 61

D2 Dopamine Receptor 2

D5 Dopamine Receptor 5

DAT Dopamine Transporter

DCM Dichloromethane

DIAD Diisopropylazodicarboxylate

DIBAL-H Diisobutylaluminum Hydride

DMP Dess-Martin Periodinane

DMS Dimethyl Sulfide

DMSO Dimethyl Sulfoxide

ECIS Electric Cell-Substrate Impedance Sensing

EDCI 1-Ethyl-3-(3-dimethylaminopropyl)carbodiimide

EGF Epidermal Growth Factor

ELISA Enzyme-Linked Immunosorbent Assay

EP Prostaglandin E Receptor

EP1 Prostaglandin E Receptor 1

EP2 Prostaglandin E Receptor 2

EP3 Prostaglandin E Receptor 3

EP4 Prostaglandin E Receptor 4

EPA Eicosapentaenoic Acid

ETA Eicosatetraenoic Acid

FAA Fatty Acid Amide

FAAH Fatty Acid Amide Hydrolase

FACS Fluorescence-Activated Cell Sorting

FLIPR Fluorometric Imaging Plate Reader Assay

FP Prostaglandin F Receptor

GDE1 Glycerophosphodiesterase 1

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GP-AEA Gylcerophospho-Anandamide

GPCR G Protein-Coupled Receptor

HCB Human Ciliary Body

h-CM Human Ciliary Muscle

HEK Human Embryonic Kidney Cells

HETE Hydroxyeicosatetraenoic Acids

HPLC-MS High Pressure Liquid Chromatography – Mass Spectrometry

h-TM Human Trabecular Mechwork

hTOB Human Transducer of ErbB2 in Osteoblast

IFNγ Interferon-Gamma

IL-1 Interleukin 1

IL-1β Interleukin 1 Beta

IL-2 Interleukin 2

IL-12 Interleukin 12

IOP Intraocular Pressure

IP3 Inositol Triphosphate

LFA Latanoprost Free Acid

LG Leaving Group

LPS Lipopolysaccharide

LTP Long-Term Potentiation

LXA4 Lipoxin A4

LXB4 Lipoxin B4

M1 Muscarinic Acetylcholine Receptor 1

mCPBA meta-Chloroperoxybenzoic Acid

MLC Myocin Light Chain Kinase

MLR Mixed Lymphocyte Reaction

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MOMCl Methoxymethyl Cloride

mRNA Messenger Ribonucleic Acid

MS Mass Spectometry

Ms Methanesulfonyl, Mesyl

MTPA α-Methoxy α-Trifluoromethylphenylacetic Acid, Mosher’s Acid

NAE N-acylethanolamide

NAPE N-arachidonoylphosphatidylethanolamide

NAPE-PLD NAPE-selective Phospholipase D

NK Natural Killer Cells

NMDA N-methyl-D-aspartate Glutamate Receptor

NMR Nuclear Magnetic Resonance

NSAID Nonsteriodal Antiinflammatory Drug

Nu Nucleophile

OEA N-Oleoylethanolamide

Ox Oxidant

pAEA Phospho-anandamide

PC Phosphatidylcholine

PDGF Platelet-Derived Growth Factor

PE Phosphatidylethanolamine

PEA N-Palmitoylethanolamide

PFC Plaque-Forming Cell Assay

PGD2 Prostaglandin D2

PGD2-EA Prostaglandin Ethanolamide D2, Prostamide D2

PGE2 Prostaglandin E2

PGE2-EA Prostaglandin Ethanolamide E2, Prostamide E2

PGF2α Prostaglandin F2α

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PGF2α-EA Prostaglandin Ethanolamide F2α, Prostamide F2α

PGG2 Prostaglandin G2

PGH2 Prostaglandin H2

PGI2 Prostaglandin I2

PGI2-EA Prostaglandin Ethanolamide I2, Prostamide I2

PLA1 Phospholipase A1

PLA2 Phospholipase A2

PLC Phospholipase C

PMN Polymorphonuclear Neutrophils

PMSF Phenylmethylsulfonyl Fluoride

PPARγ Peroxisome Proliferator-Activated Receptor

PTGFR Prostaglandin FP Receptor Gene

PTGS1 Cyclooxygenase 1 Gene

PTGS2 Cyclooxygenase 2 Gene

PTX Pertussis Toxin

Pyr Pyridine

Rf Retention Factor

RIA Radioimmunoassay

ROI Region of Interest

RT-PCR Reverse Transcription Polymerase Chain Reaction

SAR Structure-Activity Relationship

SEA N-Stearoylethanolamide

SI Suppression Index

SN2 Bimolecular nucleophilic substitution

SRBC Sheep Red Blood Cells

TBAF Tetra-n-Butylammonium Fluoride

TBDPS t-Butyldiphenylsilyl

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TBS = TBDMS t-Butyldiphenylsilyl

T-cells Thymus Cells

TES Triethylsilyl

Tf Trifluoromethanesulfonyl, Triflate

TFA Travoprost Free Acid

TGF-β1 Transforming Growth Factor Beta 1

THC Tetrahydrocannabinol

THF Tetrahydrofuran

TIPS Triisopropylsilyl

TLC Thin Layer Chromatography

TMEV Theiler’s Murine Encephalomyelitis Virus

TMS Trimethylsilyl

TNFα Tumor Necrosis Factor Alpha

Tos p-toluenesulfonyl, Tosyl

TP Thromboxane Receptor

TPA 12-O-Tetradecanoylphorbol-13-acetate

TRPV1 Transient Receptor Potential Vanilloid 1

TxA2 Thromboxane A2

UV Ultraviolet

wt Wild Type

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Chapter 1: Biosynthesis and Pharmacology of Prostaglandin Ethanolamides (Prostamides)

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Prostaglandin ethanolamides (prostamides) are endogenous lipids mediators of the eicosanoid class of compounds. Similar in structure to the prostaglandins, interest has been incited concerning prostamide’s biological actions and physiological purpose diverse from those of prostaglandin. (Figure 1.1) While their biological function has yet to be resolved, the biosynthetic pathway for the in vivo production of prostamides has been thoroughly elucidated.1-3 From the release of the fatty acid amide precursor, anandamide, to the well-

established oxidative metabolism by cyclooxygenase-2 enzyme (COX2), the presence of

prostamide at biologically significant levels has been postulated.

Figure 1.1 Two representatives of the prostamide and prostaglandin classes of eicosanoids

While prostamide research is in its infancy, mounting evidence suggests that

prostamides act in ways disparate from the prostaglandins.4-9 Studies demonstrate

prostamide exhibits an enhanced metabolic stability in vivo indicating the biological actions

of prostamide are not due to conversion to prostaglandin.8-10 Additionally, prostamides have been shown to be essentially inactive at prostaglandin receptors and to activate unique cells within the same tissue.4,7-9,11 Still, some research presents conflicting evidence suggesting that prostamides act only as prostaglandin prodrugs or prostaglandin-mimetics. These studies demonstrate that a small amount of prostamides are hydrolytically converted to prostaglandin and are sufficient to activate prostaglandin receptors.5,12-18 They claim furthermore that prostaglandin receptor gene knockout mice are unresponsive to prostamides.19,20 Examining evidence from both sides of the conflict over the distinctiveness

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Part A: Prostamide Biosynthesis

1.1 Biosynthesis, Action, and Degradation of the Prostamide Precursor, Anandamide

The biosynthetic cascade that leads to the formation of the natural prostamides begins with the release of the endogenous fatty acid amide (FAA) compound, N- arachidonoylethanolamide. A member of the N-acyl ethanolamide (NAE) class of signaling lipids, N-arachidonoylethanolamide, also known as anandamide (AEA), is produced from membrane phospholipids such as phophatidylethanolamine (PE), phosphatidylcholine (PC), and phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2) and released from the membrane by the action of various enzymes.21,22 (Figure 1.2) Three distinct pathways of AEA formation from the phospholipids have been elucidated. All of these pathways begin with

Figure 1.2 Representation of the membrane phospholipids from which the NAEs are biosynthesized

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the transfer of an arachidonoyl chain from a membrane phospholipid onto the primary

amine of PE by a calcium-dependent transacylase (Ca-TA) to form N-arachidonoyl

phosphatidylethanolamine (NAPE). (Figure 1.3) From here, one pathway involves

hydrolysis of the phosphatidyl esters by either phospholipase A1/2 (PLA1/2) or by α/β-

hydrolase-4 (ABHD4) to glycerophospho-anandamide (GP-AEA) followed by cleavage of the

glycerophosphate by glycerophosphodiesterase-1 (GDE1), producing the free AEA.23,24 A second pathway begins with cleavage of the phosphatidyl group by phospholipase C leaving the phospho-anandamide (pAEA) and follows with cleavage of the phosphate group by

phosphatase giving AEA.25,26 The third and final known pathway is more applicable to the long chain saturated and monosaturated NAEs like N-oleoyl ethanolamide (OEA), N- palmitoyl ethanolamide (PEA), and N-stearoyl ethanolamide (SEA); however AEA can also be biosynthesized in this way. This single enzymatic step involves complete hydrolysis of the phosphatidyl moiety by a NAPE-selective phospholipase D (NAPE-PLD) to give AEA.27,28

Figure 1.3 The pathways for biosynthesis of anandamide from membrane phospholipid, phosphatidylethanolamine.

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Anandamide is an agonist of the cannabinoid (CB) receptors, naturally occurring throughout the body and brain. AEA has been shown to mimic several of the pharmacological effects of the psychoactive cannabinoid agonist, tetrahydrocannabinol

(THC), including analgesia, hypothermia, and motor defects.29 The endocannabinoid signaling system has also been indicated in tissue protection against pathological insult or injury, especially within the CNS.30-33 Anandamide elicits its cannabinergic effects by acting as a partial CB1 receptor agonist with modest potency (Ki = 240 nM) and a relatively weak

34,35 CB2 receptor agonist (Ki = 0.44-1.93 µM).

Figure 1.4 Representation of the inactivation of AEA’s cannabinergic action through active uptake by a membrane transporter protein followed by hydrolysis by FAAH to AA.

The principal enzyme responsible for AEA metabolism and the termination of AEA signaling is the well-characterized, integral membrane protein, fatty acid amide hydrolase

(FAAH; EC 3.5.1.99).36 (Figure 1.4) Anchored to the interior of the cell, FAAH’s substrates may be transported from the extracellular space through a transporter protein. Due to its lipid structure, AEA may also diffuse from the phospholipid bilayer and into the FAAH active site directly from this membrane. The product of FAAH’s hydrolysis of AEA, arachidonic acid

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(AA), is inactive at the cannabinoid receptors. Thus this enzymatic transformation brings an

end to AEA’s cannabinergic actions. (see Section 5.1)

1.2 Oxidative Metabolism of Fatty Acids and Fatty Acid Amides by Cyclooxygenase Enzymes

Anandamide serves as a substrate for oxidative metabolism by the cyclooxygenase

enzyme COX-2. Cyclooxygenase (COX) is known principally as the enzyme responsible for

the oxidative metabolism of arachidonic acid into prostaglandin H2 (PGH2), the precursor for the formation of the important lipid mediators, prostaglandins and thromboxanes. These enzymes are widely studied in the treatment of inflammatory conditions, such as rheumatoid arthritis and osteoarthritis, due to the knowledge that nonsteroidal antiinflammatory drugs (NSAIDs) elicit their effects primarily through inhibition of the COX enzymes and the subsequent formation of prostaglandins.37 The mechanistic actions of COX

on arachidonic acid to produce prostaglandins can be extended to the parallel process of its

oxidative metabolism of anandamide to the prostamides.

a b S/D COOH COOH

S/D Arachidonic Acid (AA) Polyunsaturated Fatty Acid (20:4) COOH 2O2 Cyclooxygenase

Eicosapentaenoic Acid (EPA) (20:5) O COOH S/D COOH O S/D Eicosatrienoic Acid (ETA) OOH (20:3) O H dro erox endo eroxide y p y p OH N 2e- H Peroxidase Anandamide (AEA)

O COOH OH S/D O O O OH S/D OH 2-Arachidonoylglycerol (2AG) Hydroxy endoperoxide Figure 1.5 (a) General oxygenation of polyunsaturated fatty acids to hydroperoxy endoperoxides. The S/D label represents presence of either a single or double bond. (b) Substrates for the cyclooxygenase enzymes

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The primary function of COX is a double step catalysis involving first, the

oxygenation of polyunsaturated fatty acids to hydroperoxy endoperoxides and second, the

reduction of the hydroperoxide to a hydroxy endoperoxide.38 (Figure 1.5) 5,8,11,14-

Eicosatetraenoic acid (20:4), more commonly referred to as arachidonic acid (AA), is the preferred substrate for this enzymatic reaction. However other fatty acids, such as

5,8,11,14,17-eicosapentaenoic acid (EPA, 20:5), and 8,11,14-eicosatrienoic acid (ETA,

20:3), have also been shown to undergo catalysis, but at much slower rates (relative efficiencies: (20:4 > 20:3 >> 20:5).39 (Figure 1.5b) The reaction exclusively produces prostaglandin H2 (PGH2) which is then metabolized further by individual isomerases or a

40 reductase to give the prostaglandins (PGE2, PGF2α, PGD2, PGI2) and thromboxane (TxA2).

(Figure 1.6)

COOH AA

COX (cyclooxygenase)

O COOH PGG2 O

OOH COX (peroxidase)

O COOH PGH2 O

OH

Prostaglandin Synthases α ( )

HO O HO α α α α ( ) ( ) ( ) ( ) O ω ω ω ω ω ( ) ( ) ( ) ( ) O ( ) O HO HO α HO PGD2 PGE2 PGF2 PGI2 TxA2

α-c a n COOH ( h i ) ω ( -chain)

OH

Figure 1.6 The biosynthetic scheme of prostaglandins

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1.2.1 Mechanism of Fatty Acid Oxidation

1.2.1.1 Autoxidation of Fatty Acids

ox O2

H H kinetic thermodynamic

O O HO O + H H Figure 1.7 Autoxidation of polyunsaturated fatty acids

The mechanism for the conversion of arachidonic acid to the prostaglandins is comparable to that of polyunsaturated fatty acid autoxidation. Known to be involved in the rancidification of foods, autoxidation entails the oxidation of a methylene between cis double bonds to a carbon-centered radical that is then trapped by a molecule of oxygen forming a hydroperoxide.41 (Figure 1.7) The initiation of this radical reaction is the removal of a bis- allylic hydrogen by an oxidant to generate a carbon radical. Allylic resonance distributes the electron density between the central carbon and those at either end. The propagation step begins when molecular oxygen traps the radical producing a peroxyl radical that is then available to react with another fatty acid molecule forming peroxide and another carbon- centered radical. Longer chain polyunsaturated fatty acids can undergo intramolecular cyclization of the peroxyl radical intermediate to give mono- and bicyclic peroxides. These however, are formed as very minor products (~1-2%)42,43 (Figure 1.8) Included in these cyclic peroxide products is a precursor to the isoprostanes. These compounds have identical functionality to the prostaglandins, however the 4,5-dialkyl substitutions are in a cis configuration rather than the characteristic trans of the prostaglandins. The similarity of these compounds to the prostaglandins illustrates the intrinsic ease by which fatty acids can be oxidized to prostaglandin-like compounds. The role of the COX enzyme must therefore

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be to spatially and energetically promote the formation of these bicyclic endoperoxide

compounds and initiate the selective formation of the five new chiral centers.

COOH

COOH

O-O

OOH COOH COOH O O O O

OOH Figure 1.8 Examples of intramolecular cyclization during autoxidation of fatty acids

1.2.1.2 Cyclooxygenase Crystal Structure and Active Site Interactions

The cyclooxygenase enzyme is a homodimer with a molecular mass of 70 kDa per

monomer and consists of three distinct structural domains: a short N-terminal epidermal

growth factor domain, an α-helical membrane-binding motif, and a C-terminal catalytic

domain.45 (Figure 1.9) It is an integral monotopic membrane protein with the membrane-

binding motif comprised of a series of amphipathic α-helices bound to a membrane

monolayer. The catalytic domain is highly homologous to mammalian peroxidases

containing two separate but mechanistically coupled active sites and making up

approximately 80% of the protein. Two isoforms of COX have been identified containing

highly homogenous 3D structure and 60% identical amino acid sequence, but differential

expression.46 COX catalyzes two sequential reactions within the distinct active sites. The first is the oxygenation of AA to PGG2 and takes place within the cyclooxygenase active site.

The second is a reduction of the PGG2 peroxide to the PGH2 alcohol within a peroxidase active site.47

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Catalytic

Peroxidase Active Site

Epidermal growth factor- Cyclooxygenase like domain Active Site

Membrane Binding Domain Figure 1.9 Protein crystal structure of COX-144

Figure 1.10 Stereo view of AA bound in the cyclooxygenase active site channel48

The cyclooxygenase active site is located at the end of a long hydrophobic channel that is broad near the membrane binding domain and narrows as it extends inward. A

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constriction formed by Arg120, Tyr355, and Glu524 separates the cyclooxygenase active

site from the broad end of the channel.48 (Figure 1.10) This site is subdivided into a

hydrophobic main substrate binding channel and a smaller amphipathic side pocket. The

substrate binds in an extended L-shape orientation allowing the carboxylate ion to form an

ion pair with the guanidinium group of Arg120 and a hydrogen bond with Tyr355 at the

bottom of the active site.48 The ω chain is surrounded by 6 aromatic amino acids and binds

to a narrow channel at the top of the active site. The central segment adopts an S shape

around the sidechain of Ser530. This serine is integral for the creation of the 15-(S)

hydroxyl chiral center as shown by mutation studies replacing Ser530 with Met530, Thr530,

and Val530 which produce 15-(R) prostaglandins.49

1.2.1.3 Catalysis of Fatty Acid Oxidation by Cyclooxygenase

Catalytic conversion of AA by COX is initiated when a peroxide, likely a fatty acid hydroperoxide or a peroxynitrite, reduces the heme cofactor near the peroxidase active site

to a ferryl-oxo derivative.50 This derivative in turn oxidizes Tyr385 within the cyclooxygenase active site to produce a tyrosyl radical.51 The radical abstracts the 13-pro(S)

hydrogen leaving a carbon-centered radical at the C-13 position. (Figure 1.11) Migration of

the radical to C-11 forms a more thermodynamically stable pentadienyl radical and is

followed by trapping with molecular oxygen.52 The established peroxyl radical cyclizes to the

C-9 position giving an endoperoxide and pushing the radical to C-8. A second cyclization follows with C-8 attacking the C-12 position and forming the characteristic pentane ring. A

second molecule of oxygen traps the radical at the C-15 position establishing the peroxyl

radical that will abstract the hydrogen back from Tyr385 giving the prostaglandin G2 (PGG2)

intermediate and the tyrosyl radical. PGG2 peroxide subsequently migrates to the peroxidase active site where it is reduced to the alcohol product, prostaglandin H2 (PGH2),

-31- and in the process oxidizes the heme iron back to the ferryl-oxo complex, thereby initiating the next turn over. (Figure 1.12)

COOH COOH COOH H

O O O OH

Tyr 385 Tyr 385

COOH COOH COOH O O O O O O

COOH O O COOH O COOH O O O

O COOH O COOH O COOH O O O

O O O O O O O H

Tyr 385 O COOH O

HO O

O

Tyr 385

Figure 1.11 Mechanism of catalysis by cyclooxygenase

O COOH + [(PPIX)Fe3 ] O HO O

O COOH O

OH + + + [(PPIX)Fe4 O] [(PPIX ) Fe4 O] Figure 1.12 Mechanism of catalysis by peroxidase

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1.2.2 Differentiation of the Cyclooxygenase-1 and Cyclooxygenase-2 Isoforms

While the two COX isozymes have similar structure and biochemical functionality, their most notable distinction is in their expression profiles. Cyclooxygenase-1 (COX-1) is expressed constitutively and is involved in the maintenance of cellular homeostasis.53 COX-1

enzyme levels do not vary significantly regardless of cellular environment. PTGS1 is the

gene responsible for the constitutive transcription of a very stable 2.8-kb mRNA which then

encodes the COX-1 protein.54 Cyclooxygenase-2 (COX-2), conversely, is expressed on

demand in response to a number of stimuli, including inflammatory cytokines (IL-1, TNFα), growth factors (EGF, PDGF), pyrogens (lipopolysaccharide), and tumor promoters (TPA, benzo[α]pyrene).2,55-58 COX-2 is encoded by a 4-kb mRNA that undergoes rapid turnover and is transcribed by the immediate early gene, PTGS2.59 While COX-2 is also expressed

constitutively in specialized regions of the brain and kidney, it is by all accounts the more

inducible of the enzymes.

While the isoforms have similar structures (60% sequence identity), there are a

few key variances in their amino acid sequence that create substantial differences in their

binding profiles. The most notable diversity is found in the size of the cyclooxygenase active

site as the volume of the active site of COX-2 is approximately 27% larger than that of

COX-1.60 This difference results primarily from the substitution of a sterically less

demanding valine (Val-523) residue of COX-2 in place of a more bulky isoleucine (Ile-523)

of COX-1.47 Other contributing substitutions in this area include a Val-434 and Arg-513 in

COX-2 for Ile-434 and His-513 in COX-1, respectively. The result is the presence of a side

pocket near the base of the active site found exclusively in COX-2. (Figure 1.13) This

simple distinction gives rise to significant differences in the ability of the enzymes to bind

both substrates and inhibitors. Because of the larger binding pocket, COX-2 can oxygenate

larger, neutral derivatives of AA much more efficiently than COX-1. These substrates that

are selectively metabolized by COX-2 include the endocannabinoids, N-

-33- arachidonoylethanolamide (anandamide - AEA), and 2-arachidonoylglycerol (2-AG).1,62

(Figure 1.14)

Figure 1.13 Comparative stereo views of (a) AA bound to COX-1 emphasizing smaller binding site constrained by Ile523 and (b) PGH2 bound to COX-2 illustrating the larger side pocket allowed by Val52361

O

OH N H

Anandamide (AEA) N-arachidonoylethanolamide

O OH

O

OH 2-Arachidonoylglycerol (2AG) Figure 1.14 Endocannabinoid selective substrates for COX-2

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In addition to binding larger substrates, crystal structure studies have revealed that

COX-2 possesses the unique ability to bind AA and EPA in an alternate binding mode.63 In this inhibitory mode, the orientation of the substrate is opposite to the catalytic

conformation, and catalysis is not viable. As opposed to the catalytic conformation, the

carboxylate ion binds near the top of the cyclooxygenase active site forming hydrogen

bonds with Tyr-385 and Ser-530. In this reverse orientation the Arg-120 forms van der

Waals interactions with the ω tail of the fatty acid, rather than forming an ion pair with the

carboxylate ion. (Figure 1.15)

Figure 1.15 Stereo view of the inhibitory binding mode of AA unique to COX-261

As a result of having this inhibitory mode of binding, direct ligand-binding studies

show AA binds to COX-2 with an affinity (Kd) 10-100-fold higher than predicted from its

63 observed KM for AA. The inhibition of COX-2 by aspirin is also unique in that while aspirin

acetylation of COX-1 halts its oxygenase activity altogether, it merely shifts the primary

products of COX-2 from PGH2 to 15-(R)-hydroxyeicosatetraenoic acid (HETE), exclusively leading to downstream lipoxin production.64,65 (Figure 1.16) Therefore, there is sufficient

space within the active site of COX-2 to allow for both the acetylation and for partial

oxygenation of the substrate. Finally, COX-2 can be selectively inhibited by numerous

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compounds including most notably the diarylheterocyle sulfones or sulfonamides such as

rofecoxib, celecoxib, and valdecoxib. (Figure 1.18)

Tyr385

O

O O H Ser530 Ser530 H O O OH O Tyr348 O H O O- O- O

Figure 1.16 Mechanism of acetylation of COX by aspirin

1.2.3 Inhibition of Cyclooxygenase

O OH O OH O OH O O H3CO Aspirin Ibuprofen Naproxen

O O OH HO O H H CO O 3 H HN N N N N S O O O

Mefenamic acid Piroxicam

Cl Indomethacin Figure 1.17 Commonly known NSAIDs

Inhibition of the COX enzymes has long been a principal therapeutic target for the treatment of acute and chronic inflammation. Non-steroidal anti-inflammatory drugs

(NSAIDs) such as aspirin, ibuprofen, and naproxen, elicit their effects by inhibition of both

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COX enzymes, decreasing the resultant amount of pro-inflammatory prostaglandin products.

(Figure 1.17) A well-established side-effect of these nonselective therapeutics is irritation

of the gastrointestinal tract caused by the inhibition of COX-1 within the stomach mucosa.66

Because COX-2 is the isoform expressed on demand in response to inflammatory stimuli such as lipopolysaccharide (LPS) and cytokine interleukin-1 beta (IL-1β), it follows that selective inhibition of COX-2 would provide significant decrease in prostaglandin synthesis during inflammatory response without effecting the homeostatic processes of COX-1 prostaglandin production. This, in fact, is the case with the selective COX-2 inhibitors rofecoxib (Vioxx), celecoxib (Celebrex), and valdecoxib (Bextra), also known as the coxibs, which have been shown to successfully decrease inflammation with a drastic reduction in the ulcerogenic effects associated with the NSAIDs.67-69 (Figure 1.18)

O O O O O O S S S H2N H2N

N N O CF3 O N O

Rofecoxib Celecoxib Valdecoxib Figure 1.18 The coxibs, selective COX-2 inhibitors

The coxibs, while effectively treating inflammation with less gastrointestinal toxicity,

give rise to a serious cardiovascular toxicity that had not been predicted from the

gastrointestinal clinical trials lasting one year.70,71 In addition, studies of COX-2 knockout animals showed no signs that the absence of COX-2 activity leads to an increased risk of cardiovascular events such as myocardial infarction.72,73 However, when rofecoxib and celecoxib underwent clinical trials for use in colon adenoma prevention over a three year period, both drugs saw a two to three fold increase in cardiovascular side effects including myocardial infarctions and ischemic cerebrovascular events.74,75

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This cardiovascular toxicity can be explained by consideration of the reduction of

prostacyclin (PGI2) concentration relative to that of thromboxane A2 (TxA2). Most tissues convert PGH2 to a few of the final prostaglandin and thromboxane products, but not all five.

For example, prostaglandin synthesis in vascular endothelium produces mainly PGI2 and

PGE2 whereas those in blood platelets principally produce TxA2 and PGD2. (Figure 1.19)

AA

COX-1 COX-2

PGH PGH2 PGH2

TXS PGDS PGFS PGIS cPGES mPGES

PGF PGI PGE TXA2 PGD2 PGF2a PGI2 PGE2 Figure 1.19 COX-1 versus COX-2 prostaglandin biosynthesis cascade

PGI2 is a potent vasodialator and reduces the responsiveness of platelets to proaggregatory substances, while TxA2 is itself proaggregatory, increasing platelet aggregation and stimulating activation of new platelets. Because COX-1 is associated with platelets and COX-2 can be induced in vascular endothelial cells, selective inhibition of COX-

2 decreases prostacyclin as thromboxane levels remain unchanged. Chronic use of selective

COX-2 inhibitors leads to a substantial imbalance of prostacyclin to thromboxane and increases the chance of platelet aggregation leading to the formation of harmful clots and cardiovascular events. (Figure 1.20) This phenomenon is not observed with NSAIDs due to

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their ability to inhibit both COX enzymes and therefore decrease the production of

prostacyclin and thromboxane evenly.

Figure 1.20 Illustration of blood clotting homeostasis. Platelets produce proaggregatory TXA2 while 76 endothelial cells produce vasodilator PGI2

In addition to being selective for certain inhibitors, COX-2 can also selectively recognize the neutral arachidonic acid derivatives anandamide (AEA) and 2- arachidonoylglycerol (2-AG). The metabolism of these endocannabinoid compounds leads to a set of endogenous compounds analogous to the prostaglandins.1,62 The thromboxanes are

the exception, as PGH2-EA and PGH2-G are poor substrates for thromboxane synthase. This raises the possibility that inhibitors of COX-2 may exhibit their pharmacological effects not only through reduction of the prostaglandins, but also through reduction of the prostamides.

Part B: Prostamide Pharmacology

The biological significance of the prostamides has yet to be established, and great

contention has been made over the novelty associated with prostamide analog therapeutics.

The essential question posed is whether prostamide compounds exhibit their therapeutic

actions through activation of the well-established prostaglandin receptors or through

selective activation of one or several yet unidentified “prostamide receptors”. A plethora of

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evidence suggests that prostamides indeed act on receptors divergent of the classical

prostaglandin receptors.4-9 Studies that support this hypothesis find that prostamides are metabolically stable and will not hydrolyze to prostaglandins in vivo,8-10 are essentially

inactive at prostaglandin receptors,4,7-9,11 exhibit activity in tissue that is much higher than

expected based on isolated prostaglandin receptors,9 activate entirely different cells than the prostaglandins,7 are blocked by selective antagonists that do not affect prostaglandin

agonism,5,8 exhibit activity in T-cells lacking FP receptors,77 exhibit no activity in osteoblasts

cells containing FP receptor gene,77 and successfully treat glaucoma in patients that are unresponsive to prostaglandin therapies.78 Still, some investigators insist that, at least in the case of glaucoma therapeutics, the biological purpose of prostamides is simply to act as prostaglandin prodrugs and/or prostaglandin-mimetics. Studies that maintain this stance

find that a small but essential amount of prostamides do hydrolyze in vivo,5,12-18 the

concentration of prostamide after topical dosing is more than enough to overcome a low

activity at prostaglandin receptors,6,79,80 tissues exhibiting high activation by prostamides

contain super-sensitive and super-coupled prostaglandin receptors,80 FP prostaglandin

receptor gene knockout mice are unresponsive to prostamides,19,20 and the observed pharmacology of prostamide and prostaglandin glaucoma treatments are identical.80,81

Currently, prostaglandins and prostamides have found great therapeutic usefulness

in treating ocular hypertension conditions such as glaucoma, and it is within the realm of

ophthalmology that the debate surrounding prostamides mainly focuses. Thus prostaglandin

F2α and prostamide F2α, the preferred prostanoid analogs for the treatment of glaucoma and the core configuration for glaucoma drugs latanoprost and bimatoprost, have undergone the most extensive comparative study. (Figure 1.21) Investigating the pharmacology of bimatoprost and other prostaglandin F2α glaucoma therapeutics may lead to a better

understanding of general prostamide biology. Some findings based on differences between

prostamide F2α and prostaglandin F2α can be extrapolated to the other prostamides and to

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HO HO H N COOH OH O HO HO OH OH Prostaglandin F2a Prostamide F2a

HO HO H HO N O O

O O O O HO HO HO OH OH OH

CF3 Bimatoprost Latanoprost Travoprost

Figure 1.21 Representatives of the prostanoid class of glaucoma therapeutics. Bimatoprost,

latanoprost, and travoprost are analogs based on the structures of prostaglandin F2α and prostamide

F2α.

biological systems outside the ocular system. However, caution must be taken that

ophthalmology studies of this small set of prostaglandin F2α analogs alone does not

encompass all of prostamides’ possible biological actions. This area is simply where the

most work has been done, and investigations of other prostamides in additional tissues

should be undertaken to garner a greater understanding of prostamide biology.

Controversy exists over the hydrolysis of bimatoprost, and it is complicated by

conflicting data and possibly ulterior motives. Therefore caution must be exercised when

reviewing the numerous publications on the subject of prostamide therapeutics. Most of the

ophthalmology research that follows was done by members of laboratories within the

pharmaceutical companies, Allergan Inc or Alcon Research Ltd or by collaborators with said

groups. Allergan Inc, designers of Lumigan® (bimatoprost ophthalmic solution) and Latisse

desire their compound be uniquely active and distinct from the prostaglandin analogs such

as latanoprost and travoprost. As such, their research adamantly supports the hypothesis

that prostamides such as bimatoprost are extremely stable and do not undergo hydrolysis

to prostaglandins and that their actions are disparate to those of prostaglandin receptor

agonists.4-8,10,11,13,79,82-85 Extensive evidence of prostamides’ diverse actions will be discussed herein. Taking up the opposing position, Alcon Research Ltd is the company responsible for

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the development and marketing of the prostaglandin pro-drug Travatan Z® (travoprost

ophthalmic solution). The research coming from this laboratory exclusively maintains the

position that bimatoprost is simply a pro-drug and behaves just like every other prostaglandin analog anti-glaucoma therapeutic.14-17,80,86-88 By negating the novelty of

bimatoprost’s method of action Alcon stands to equate their own product’s abilities with that

of bimatoprost. It is evident that both companies have a stake in whether their drugs are

seen as prodrugs with the action of classical prostaglandins or whether they represent a

novel class of ocular therapeutics. Therefore caution must be used when interpreting any

data given by these groups. Regardless of whether bimatoprost’s actions are unique, it has

become the most widely used and effective singular anti-glaucoma therapeutic currently on

the market.

1.3 Debate Concerning Hydrolysis of Prostamides to Prostaglandins

The first critical point of contention involves metabolism of the prostamides and

whether hydrolysis of the amide bond to reveal the free acid prostaglandin is necessary for

prostamides to give biological activity. In addition, the amount of FAAH hydrolysis to free

acid observed in vivo and whether the level of resulting free acid metabolite is sufficient to

activate prostaglandin receptors is under debate.

1.3.1 Evidence of Negligible Prostamide Hydrolysis

Studies show that after four hour incubation in rat brain, lung, liver and in feline and

human iris-ciliary body homogenates, the prostamides exhibit extremely low conversion by

hydrolysis to their free acids.8 This level of hydrolysis is considered to be negligible,

especially when compared with the hydrolysis of the endocannabinoid anandamide to the

cannabinoid-inactive arachidonic acid. (Table 1.1) Fatty acid amide hydrolase (FAAH) is the principle enzyme responsible for this conversion, and FAAH metabolism in fact is the major pathway of deactivation for anandamide in endocannabinoid signaling.89-91

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Table 1.1 Degree of hydrolysis of the prostamides versus anandamide after four hour incubation in rat tissue homogenates.8 % hydrolysis to % hydrolysis to % hydrolysis to free acid in rat free acid in rat free acid in rat brain lung liver

PGE2-EA 0.1 - 2.6 - -

PGD2-EA 1.6 - 3.6 - -

PGF2α-EA 0.2 - 0.6 0 - 1.7 2.2 - 2.9

Anandamide 87.2 - 96.7 46.7 - 49.3 85.3 - 100

Evaluation of the metabolism of prostamide drug bimatoprost and prostaglandin

ester prodrug latanoprost to their free acids in human iris-ciliary body tissue homogenates

reveals markedly different hydrolysis rates.79 (Figure 1.22) While the total concentration of bimatoprost remains relatively unchanged, there is no evidence of an increase of bimatoprost free acid metabolite over a three hour incubation period. Latanoprost

conversely shows almost complete conversion to latanoprost free acid within one hour of

incubation and its status as a prodrug is confirmed. Because no measurable free acid

metabolite of bimatoprost is detected, bimatoprost cannot be considered a prodrug when

eliciting its effects on intraocular pressure.

Figure 1.22 Conversion of bimatoprost (a) and latanoprost (b) to the free acid metabolite in human iris-ciliary body tissue homogenates.79 (a) Total bimatoprost concentration (solid triangle), bimatoprost free acid metabolite (open triangle). (b) Total latanoprost concentration (solid square), latanoprost ester (solid circle), latanoprost free acid metabolite (open circle).79

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a b

Figure 1.23 (a) Lack of inhibitory effect of prostamides E2, F2α, D2 and bimatoprost (50µM and 100µM) on FAAH enzymatic hydrolysis of [14C]anandamide activity.8 (b) Lack of increase in displacement of [3H]CP55940 binding to CB receptors by 100nM anandamide in the presence of 9 prostamide E2. FAAH inhibitors PMSF and OL-093 are positive controls.

The stability of prostamide to FAAH hydrolysis has been confirmed by studies in which prostamide is found not to be a substrate/inhibitor of FAAH.8,9 In a direct study, it

was shown that prostamides E2, D2, and F2α and bimatoprost do not inhibit FAAH’s hydrolysis of radiolabeled anandamide at 50 µM and 100 µM (Figure 1.23a).8 This inability

to bind FAAH was confirmed indirectly by studies measuring the amount of specific binding

of anandamide to its CB1 receptor in the presence of FAAH and PGE2-EA and by studies

administering FAAH inhibitor along with prostamide in tissue assays.9 Anandamide’s ability to displace radiolabeled CB1 agonist [3H]CP-55940 from CB1 is unaffected by the addition of

PGE2-EA. Meanwhile as expected, administration of FAAH inhibitors PMSF and OL-093

greatly enhances displacement (Figure 1.23b). This strongly suggests that PGE2-EA is not

a FAAH inhibitor nor a FAAH substrate as it does not slow the inactivation of anandamide

CB1 signaling. In an analogous study of the guinea pig vas deferens preparation, known to

9 contain EP3 receptors, the presence of PMSF does not alter the potency of PGE2-EA. The

result indicates that PGE2-EA action at EP3 is not the result of hydrolysis to PGE2 prior to EP3 receptor activation.

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The tiny amount of bimatoprost free acid is insufficient to act on prostaglandin FP

receptor, and therefore bimatoprost action cannot be due to a necessary conversion to

prostaglandins.6,10 In fact, in an exaggerated study using a dose concentration 80 times the

Cmax of bimatoprost present within the cornea after a single 0.1% dose, the hydrolysis rate

was only 0.4% per hour over 23 hours.92 A 0.1% percent dose equates to three times the

therapeutic dose and gives a maximum corneal concentration of 1.5 µM after a single dose

and 7.0µM after multiple doses.6 Employing 120 µM of bimatoprost directly to the cornea

and incubating for an especially large amount of time (24 hours), much longer than

bimatoprost is present at the corneal surface, gave only 10% total conversion to the free acid. (Figure 1.24) To be considered a prodrug like latanoprost, a rapid and complete hydrolysis to an active metabolite would be expected. As this is clearly not the case with bimatoprost, its therapeutic actions are probably the result of the intact amide and not through hydrolysis to the free acid.

Figure 1.24 HPLC chromatogram of a sample of human cornea incubated with 120µM bimatoprost for 23 hours (lower trace). The upper trace is co-injection of bimatoprost and the bimatoprost free acid 92 metabolite, 17-phenyl-trinor PGF2α.

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1.3.2 Evidence of Substantial Prostamide Hydrolysis

Alternative studies refute the notion that prostamides do not undergo meaningful hydrolysis to the free acid prostaglandins.80,87,93 While the prostamide are indeed significantly more stable to hydrolysis than anandamide and the prostaglandin esters, the amount of free acids observed following incubation is not zero. In fact, the concentration of bimatoprost free acid is simply below the level of detection (25 nM) employed by the studies discussed previously.8,9 Subsequent studies have established concentrations of bimatoprost

free acid both above and below this level.12-14 One such study determined the maximum

concentration of bimatoprost free acid with repeated daily dosing (0.03%) in human

aqueous humor to be approximately 30 nM (Figure 1.25). The bimatoprost amide parent

compound on the other hand, reaches maximal levels of only about 3 nM over the five hour

testing period.14 While continuous daily dosing is necessary to reach this level of

bimatoprost free acid, it is clear that there is a significant build up of the hydrolyzed product

over time such that activation FP receptors is probable.

a b

Figure 1.25 Concentrations in individual patient aqueous humor of (a) bimatoprost free acid (17- 14 phenyl-trinor-PGF2α) and (b) bimatoprost.

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Prostaglandin compounds with their free acids have a much higher affinity for prostaglandin receptors than the amides and esters such that even small conversion of prostamide to prostaglandin in vivo would be sufficient to fully activate prostaglandin receptors. The FP receptor agonist potency values of prostaglandin F2α, bimatoprost and its free acid, and latanoprost and its free acid are given in Table 1.2. In all human ciliary tissue assays, the free acids of bimatoprost and latanoprost are considerably more potent.6,15-17 Therefore much smaller amounts of the free acid are required to activate the

FP receptors in these tissues. Essentially any concentration above approximately 4 nM within the human ciliary body will be quite sufficient to act as an agonist at the FP receptors. Therefore, even a miniscule amount of hydrolysis could mean potent activity at these receptors. According to these studies, bimatoprost free acid reaches levels ranging from 5 nM to 35 nM, well above the amount required for substantial agonism of the FP receptors within the ciliary tissue.12-14

Table 1.2 Equilibrium functional FP receptor agonist potency values (EC50) of given prostanoids and their free acids in human and feline ocular cells. Human Human Ciliary Human Ciliary Trabecular Feline Iris EC50 Body EC50 Muscle Cells 6,15 16,17 16 Meshwork Cells (nM) (nM) EC50 (nM) 16 EC50 (nM)

6 15 PGF2α 29 104 62 58 ; 18.6

Bimatoprost 681 9600 3245 346; 14015

Bimatoprost 3.3 3.8 26 0.9915 Free Acid

Latanoprost 173 313 564 15306; 11.715

Latanoprost Free 45.7 124 35 666; 29.915 Acid

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Significant discrepancy has been found within the feline iris assay depending on the

laboratory in which the testing was done, so both of the conflicting values are included in

Table 1.2.6,15 Woodward’s laboratory at Allergan, Inc. has described bimatoprost as much

more potent in the feline iris assay than the values found by Sharif’s laboratory at Alcon

Research, Ltd. While Woodward did not report on the potency of bimatoprost free acid, due

presumably to the conviction described previously that bimatoprost dose not undergo

hydrolysis, Sharif’s data suggests that bimatoprost free acid is also substantially more

potent in feline iris tissue than the parent bimatoprost compound.

Figure 1.26 shows a compilation of the observed ability of the free acids of bimatoprost (BFA), latanoprost (LFA), and travoprost (TFA) to activate FP in human ciliary body tissue, human ciliary muscle cells, human trabecular meshwork cells, and feline iris

14 tissue. By taking a ratio of the Cmax for the free acid in the tissue over the EC50 value of the free acid at the FP receptor, an expected threshold for activation can be established. Where the ratio exceeds 1, agonism of more than 50% of the FP receptors in the tissue is expected by the given free acid. The results of five different studies are compiled and in all three of the studies that investigated bimatoprost free acid, it is evident that the concentration of

Figure 1.26 Cmax/EC50 ratios for free acids of bimatoprost, latanoprost, and travoprost in human aqueous humor from studies by Alcon,14 Camras,12 Cantor,13 Calissendorf,94 Sjoquist.95

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free acid is more than enough to be active at the FP receptors in human ciliary body tissue,

human ciliary muscle cells, and feline iris tissue, but not in trabecular meshwork cells. In

fact the free acid concentration to EC50 ratio was substantially higher with bimatoprost free

acid than with either latanoprost or travoprost free acids. This lends outstanding support to

the hypothesis that bimatoprost is sufficiently metabolized to the free acid to elicit its

hypotensive effects within the ocular system solely through activation of FP receptors.

Clinical studies have shown that even a singular dose of bimatoprost causes

significant reduction of intraocular pressure.18 A study investigating the concentration of bimatoprost free acid in aqueous humor following a single dose thus proves useful. For while an accumulation of bimatoprost free acid occurs with daily dosing over long periods of time, the increased amount of free acid still may not be responsible for the actions of bimatoprost. The average concentration of bimatoprost acid over the initial three hours post singular dosing was approximately 5.9 nM, while for the bimatoprost it was about 4.5 nM.13

(Figure 1.27) With this evidence of relatively equal portions of bimatoprost and its hydrolyzed free acid and the considerably higher potency of bimatoprost acid shown previously, clearly the intraocular effects of bimatoprost should be attributed to its hydrolysis product.

Figure 1.27 Levels of (a) latanoprost free acid (open circle) and bimatoprost free acid (solid circle) and (b) intact latanoprost ester and bimatoprost amide in samples of patient aqueous humor after a single dose.13

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In addition to unsatisfactorily high detection limits, the absence of detectible bimatoprost free acid in the assay tissue preparations may in fact indicate that the tissue under investigation acts as a sink for bimatoprost free acid due to its high affinity and slow off-rate from the large number of FP receptors. The shear density of FP receptors and the extremely high affinity of the bimatoprost acid for the FP receptor in these tissues may obscure the observed concentration of bimatoprost acid that is free in solution. Conceivably, there is an even higher concentration of bimatoprost free acid than what has been measured in the aqueous humor.

As will be discussed, prostamides have substantially less potency than their prostaglandin counterparts in most tissues, while in certain tissues, such as feline iris, they exhibit very similar potency. This phenomenon may be due to tissue specific hydrolysis by

FAAH. Tissues that tend to express higher levels of FAAH will theoretically exhibit more hydrolysis of prostamides to prostaglandins, and the resulting sensitivity of these tissues to prostamide will be very similar to that of prostaglandin. Conversely, tissues that exhibit lower expression of FAAH will be far more responsive to the prostaglandins than to prostamides.

Finally, the fact that bimatoprost is undeniably more stable to hydrolysis than the prostaglandin esters latanoprost and travoprost perhaps is the reason a significantly higher dose of active ingredient is used in bimatoprost’s drug formulation (0.03% bimatoprost vs

0.005% latanoprost and 0.004% travoprost). Assuming that all three compounds are in fact prodrugs, a compound that is hydrolyzed much less that the other two is going to need a higher dose to achieve the same effect. The fact that almost ten times as much bimatoprost is required in the formulation than latanoprost or travoprost gives weight to this theory.

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1.4 Debate Concerning Distinction Between the Biological Actions of Prostamides and Prostaglandins

The second essential point of contention involves the ability of prostamides themselves to act on the prostaglandin receptors. If prostamides exhibit sufficient potency at prostaglandin receptors, there is little need for hydrolysis in order to elicit an effect.

Assuming a similar potency to prostaglandins, this would also provide a satisfactory biological role for prostamides as prostaglandin-mimetics. While it has been well established that hydrolyzed prostamide product has the ability to potently bind the prostaglandin receptor, only with the development of prostamide glaucoma therapeutics has prostamide’s ability or lack thereof to act on prostaglandin receptors been studied.

1.4.1 Evidence of Prostamide Biological Actions Distinct from those of Prostaglandin

It has been theorized that there exists a receptor population diverse from the known prostaglandin receptors that preferentially recognizes the prostamides. Based on observed pharmacology there are two possible hypotheses – first, that certain tissues co-express prostaglandin and prostamide receptors and second, that prostaglandin receptors contain a subclass that equally recognize both prostaglandins and prostamides. Either possibility requires the identification of a yet unknown receptor to selectively bind prostamides over prostaglandins. Several studies have shown that prostamides do not meaningfully interact with any known prostaglandin receptors, meaning their half maximal inhibitory

7-9,11 concentration (IC50) is greater than 10 µM.

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1.4.1.1 Isolated Receptor and Functional Tissue Studies

Table 1.3 summarizes the findings in one extensive study and illustrates the lack of potency exhibited by the prostamides at their corresponding prostaglandin receptors.8 In both recombinant prostaglandin receptors and isolated tissue preparations known to express prostaglandin receptors, prostamides are substantially less potent at the prostaglandin receptors than their complimentary prostaglandins. With few exceptions, it is expected that concentrations of prostamides in vivo would never reach a level high enough to activate these receptors. In a Ca2+ mobilization fluorometric imaging plate reader assay (FLIPR) of

human recombinant prostaglandin receptors, prostamides exhibit affinities in the range 410-

fold to over 20,000-fold lower than their respective prostaglandins. Likewise, a radioligand

competitive binding assay on the same human recombinant prostaglandin receptors

revealed a difference of 41-fold to 2273-fold in favor of prostaglandin potency.8

Table 1.3 Equilibrium functional agonist potency values (EC50) for prostaglandins and prostamides at their respective prostaglandin receptors.8

8 8 Prostaglandins EC50 Prostamides EC50 Fold difference

Ca2+ mobilization (FLIPR) in human recombinant prostanoid receptors

PGF2α on FP 5 nM PGF2α-EA on FP >10 µM >2000

PGD2 on DP 12 nM PGD2-EA on DP >10 µM >833

PGE2 on EP1 0.2 nM PGE2-EA on EP1 848 nM 4240

On EP2 2.5 nM On EP2 >10 µM >4000

On EP3 0.3 nM On EP3 123 nM 410

On EP4 0.5 nM On EP4 >10 µM >20000

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Table 1.3 (cont’d)

Radioligand competition binding in human recombinant prostanoid receptors

PGF2α on FP 52 nM PGF2α-EA on FP 2.15 µM 41

PGE2 on EP1 13 nM PGE2-EA on EP1 10 µM 769

On EP2 27 nM On EP2 >10 µM >370

On EP3 4.4 nM On EP3 10 µM 2273

On EP4 4.8 nM On EP4 2.13 µM 444

Relaxation of intact rabbit jugular vein smooth muscle tissue (FP and EP4)

PGF2α 2.8 nM PGF2α-EA 2.0 µM 714

PGD2 28 nM PGD2-EA 3.06 µM 109

9 9 PGE2 0.45 nM PGE2-EA 89.1 nM 198

Contraction of feline iris sphincter smooth muscle (putative prostamide receptor)

PGF2α 11 nM PGF2α-EA 57 nM 5.2

PGD2 150 nM PGD2-EA 499 nM 3.3

PGE2 260 nM PGE2-EA 564 nM 2.2

Ca2+ mobilization (FLIPR) in feline recombinant FP receptor

PGF2α 6.8 nM PGF2α-EA >10 µM >1471

PGD2 40 nM PGD2-EA >10 µM >250

PGE2 396 nM PGE2-EA >10 µM >25.3

Tissue preparations that express high levels of FP receptors, including gerbil colon, mouse uterus, rat uterus, human uterus, and rabbit jugular vein have been shown to be essentially insensitive to treatment with prostamides.4 The results for the rabbit jugular vein are included in Table 1.3 and demonstrate the 109-fold to 714-fold lower affinity of the prostamides in this tissue compared to that of the prostaglandins.8

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Interestingly, there are a few tissue preparations that have been shown to equally

recognize prostaglandins and prostamides. These include feline lung parenchyma,6 rabbit uterus,84 and most notably feline iris.8,79 Table 1.3 gives the potency of prostaglandins and

prostamide with the feline iris tissue and a difference of only a small margin, 2.2-fold to

5.2-fold in favor of prostaglandin affinity, is present. This enhanced potency of prostamides

observed within the feline iris sphincter suggests that there may be receptors co-expressed

in this tissue that selectively recognize the prostamides.

In order to assess whether this phenomenon is species specific, a FLIPR study of

feline recombinant prostaglandin receptors was done to compare with the human FLIPR

study.8 The large affinity difference ranged from over 25-fold to over 1471-fold in close

agreement with the human study and in confirmation of the species-independent event. In

addition, gene regulation studies undertaken by a different lab suggest that prostamide

activity is species independent.96

The potencies of the prostamides at the recombinant prostanoid receptors are

considerably low enough to suggest that they are in fact inactive at these receptors. The

existence of a far more potent putative prostamide receptor seems like an attractive

alternative, and thus any small activity at the prostaglandins may be considered off-site

activity. Prostamide activity at wild-type and recombinant prostanoid receptors is residual

and occurs only at a concentration above 1 µM, a concentration that is unlikely to occur in

vivo.6,8,84,88,97

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a b

c d

3 Figure 1.28 Displacement of [ H]PGE2 in human recombinant prostaglandin E2 receptors: (a) EP1 (b)

EP2 (c) EP3 (d) EP4 by increasing concentrations of prostaglandin E2 (solid circles) and prostamide E2 (open circles).9

A similar study, independent of the glaucoma therapeutics debate, was done

9 focusing mainly on the major biosynthetic prostamide, PGE2-EA. Radioligand binding assays were done using membranes from cells stably transfected with EP1, EP2, EP3, and EP4 to compare the affinity of PGE2 and PGE2-EA at these receptors. In addition, PGE2 and PGE2-EA underwent functional assays using tissue preparations known to constitutively express each of these receptors. All of these assays confirm that the prostamide PGE2-EA has a much lower affinity for the prostaglandin EP receptors.9 (Figures 1.28 and 1.29)

What is interesting to note however, is the magnitude of divergence between the prostamide E2 and the prostaglandin E2 in the radioligand studies and the functional tissue

studies. The radioligand assays established a 440 to 650-fold difference in potency between

prostaglandin and prostamide whereas the functional assays showed only a 10 to 200-fold

distinction (Table 1.4).9

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a b

c d

Figure 1.29 Response of tissues expressing prostaglandin receptors as (a) contraction of guinea pig

trachea (EP1) (b) relaxation of guinea pig trachea (EP2) (c) inhibition of electrically stimulated

contraction of guinea pig vas deferens (EP3) and (d) relaxation of rabbit jugular (EP4) to increasing

concentrations of prostaglandin E2 (solid circles) and prostamide E2 (open circles). Squares in (a)

represent administration of prostaglandin E2 (solid squares) and prostamide E2 (open squares) in the 9 presence of 10 µM EP1 antagonist SC-51089.

Table 1.4 Comparison of prostaglandin E2 and prostamide E2 potencies in isolated receptor radioligand binding assays vs functional tissue assays.9 PGE EC 9 PGE -EA EC 9 Fold difference 2 50 2 50 (nM) (nM) (approx)

Radioligand binding assays hEP1 4.90 2455 500 hEP2 0.93 468 500 hEP3 0.46 200 440 hEP4 0.79 513 650

Functional tissue assays

EP1 (guinea pig trachea) 100 1000 10

EP2 (guinea pig trachea) 5.19 76.9 15

EP3 (guinea pig vas deferens) 0.82 37 45

EP4 (rabbit jugular vein) 0.447 89.1 200

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This higher potency of prostamide observed in tissues relative to prostaglandin

potency may be the result of several phenomena including species differences between

human and animal receptors, interactions of prostamides at prostaglandin receptors other

than EP receptors, interactions at other yet unknown receptors that specifically bind

prostaglandin ethanolamide and coexist with prostaglandin receptors, or resistance of

prostamide to metabolism by enzymes within the tissue that rapidly inactivate PGE2. Further

study of the interactions of prostamides diverse from those of the prostaglandins is

necessary to confirm these hypotheses and to grasp a further understanding of prostamides’

biological role.

1.4.1.2 Confocal Microscopy Studies

An exceedingly persuasive argument for the existence of a novel set of prostamide-

selective receptors was provided by a study using fluorescent confocal microscopy to

2+ observe the effects of bimatoprost, PGF2α, and 17-phenyl PGF2α on calcium (Ca ) signaling.

The results reveal that the prostamide bimatoprost consistently stimulates entirely different

7 cells from those stimulated by the prostaglandins PGF2α and 17-phenyl PGF2α.

The study entailed monitoring Ca2+ mobilization in cat iris sphincter cells using Fluo-4

dye as a fluorescent indicator. Cat iris sphincter cells were chosen due to the previously

8 discussed similarity of bimatoprost and PGF2α potency within this tissue. Cells that responded by fluorescing when treated with 100 nM or 1 µM bimatoprost, PGF2α, or 17-

phenyl PGF2α under rapid superfusion conditions were identified (Figure 1.30) and a

quantitative region-of-interest (ROI) analysis was performed (Figure 1.31). Figure 1.30

illustrates the identification of diverse cells that responded to bimatoprost (1) and PGF2α

(2,3). Bimatoprost and PGF2α activated only these disparate cells in a consistently selective

fashion.

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Figure 1.30 Image of fluorescing cells in feline iris tissue under the confocal microscope. (a) The tissue in buffered medium (b) the tissue after treatment with 100 nM bimatoprost – cell labeled 1 showed positive response (c) the tissue after treatment with 100 nM prostaglandin F2α – cells labeled 2 and 3 showed positive responses.7

17-phenyl PGF2α also excited cells 2 and 3, indicating that the C-1 amide

substitution, and not the 17-phenyl moiety, is responsible for this diverse pharmacological

response. In essence this study confirms that there is no overlap in the pharmacological

activity of bimatoprost and PGF2α and suggests that the contractive properties of the

prostamide bimatoprost on cat iris sphincter is the result of interaction with receptors

distinct from the FP prostaglandin receptors.7

To ensure desensitization or sensitization was not occurring, the sequential order in which the test substances were perfused into and aspirated from the test chamber was randomized within each set of experiments.7 A selection of the quantitative ROI results is shown in Figures 1.31 and 1.32. The randomization does not affect the resulting levels of fluorescent response as there does not seem to be any desensitization or sensitization. In addition, the bimatoprost-sensitive cells are consistently selective for bimatoprost and Ca2+ signaling was never induced by PGF2α or 17-phenyl PGF2α. The same is true for the PGF2α-

sensitive cells that cannot be acted on by bimatoprost.7

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Figure 1.31 Fluorescent Ca2+ recordings in quantitative ROI analysis induced in feline iris tissue upon

challenge with bimatoprost and PGF2α. The upper panels correspond to the cell labeled 1 in Figure 1.29b and the lower panels correspond to cell 2 in Figure 1.29c.7

Interestingly, serial confocal optical sectioning of the ROIs in this study revealed that

cells sensitive to bimatoprost have a different overall morphology than those sensitive to

PGF2a. Bimatoprost-sensitive cells were found to have a rounded, spheroidal shape while

PGF2a-sensitive cells tended to have a more elongated structure common in smooth muscle

cells.7 This again emphasizes the diversity of pharmacological action between prostamides and the prostaglandins.

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Figure 1.32 Fluorescent Ca2+ recordings in quantitative ROI analysis induced in feline iris tissue upon challenge with bimatoprost, PGF2α, and 17-phenyl-trinor PGF2α. The upper panels correspond to the cell labeled 1 in Figure 1.29b and the lower panels correspond to cell 2 in Figure 1.29c. 17-phenyl- 7 trinor PGF2α is activated only by the same cell that activates PGF2α.

1.4.1.3 Prostamide Selective Antagonists

The founding of a novel receptor requires the identification of a selective antagonist

for said receptor. A series of antagonists selective for the putative prostamide receptor has

been designed by Woodward’s lab at Allergan Inc (Figure 1.33).5,82 These proposed

antagonists do not antagonize the effects of PGF2a and its free acid analogs in tissue but instead only antagonize the contractile responses of bimatoprost and PGF2a-EA. Thus the compounds AGN-204396, AGN-204397, AGN-211334, and AGN-211335 elicit no

measurable activity at FP receptors constitutively expressed in feline iris tissue. The

establishment of these selective antagonists for the putative prostamide receptor gives yet

more weight to the proposal that prostamides do indeed have unique interactions in vivo.

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Figure 1.33 Antagonists selective for the putative prostamide receptor (a-d) and the TP receptor antagonist structures they are based on (e and f).

Development of the prototypical prostamide receptor antagonist began with the design of analogs based on the structures of TP receptor antagonists like SQ-29548 and

BMS-180291 (Figure 1.33).5 By essentially converting these compounds to ethanolamides, the initial series of selective prostamides was born. AGN-204396 and AGN-204397 are the first compounds synthesized that successfully showed selectivity for the putative prostamide receptor based on feline iris tissue assays and establish a new model for prostamide selective compounds. AGN-204396 is a concentration-dependent competitive antagonist at

5 the putative prostamide receptor (pA2 = 5.64) (Figure 1.34a).

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Figure 1.34 Effects of AGN-204396 (30 µM) on contraction of feline iris by (a) prostamide F2α, (b) 2+ 5 prostaglandin F2α, and on (c) Ca signaling in human recombinant FP receptors by prostaglandin F2α.

No antagonism of PGF2α -induced prostaglandin FP receptor contraction occurs with

the administration of ANG-204396 in the feline iris assay, and similarly no antagonism

resulted during Ca2+ signaling assay in human recombinant FP receptors (Figures 1.34b and 1.34c).5 The prostamide selective antagonist does not antagonize the effects of 17- phenyl PGF2α, verifying that the amide substitution at the C-1 head position is responsible

for this selectivity and not the 17-phenyl tail moiety. The activity of neither latanoprost free

acid nor fluprostenol is antagonized by AGN-204396, affirming the unique pharmacology of

bimatoprost. In addition, this compound antagonizes the contraction of the cat iris sphincter

by prostamide D2 and prostamide E2, but exhibites only very slight antagonism of PGD2 and

PGE2. (Figure 1.35) AGN-204395 does not antagonize the contractile effects of PGE2-G on

cat iris sphincter suggesting the prostamides and prostaglandin glycerol esters have distinct

pharmacology.

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Figure 1.35 Effects of AGN-204396 (30 µM) on the contraction of feline iris in response to (a) 5 prostamide D2, (b) prostamide E2, (c) prostaglandin D2, and (d) prostaglandin E2.

Thus it appears that at least within the confines of the cat iris sphincter assay, the

prostamides PGD2-EA, PGE2-EA, and PGF2α-EA exhibit a different agonist profile than the

prostaglandins PGD2, PGE2, and PGF2α. Because an antagonist has been developed to

selectively block the actions of only the prostamides, there must be a target for these prostamide agonists diverse from the known prostaglandin receptors.

In order to determine whether the same can be said for a selective prostaglandin antagonist which blocks only prostaglandin agonism and not that of prostamides, an FP receptor antagonist needed study. AL-8810 which has been accepted at an FP receptor antagonist/partial agonist86 was tested against the putative prostamide receptor activation and surprisingly found to be a weak but full agonist. In response, AGN-204395 was used in an attempt to block the agonism caused by AL-8810, but there was no change in activity.

(Figure 1.36) It therefore appears that the agonism by AL-8810 acts in a similar fashion to

PGF2α and, like PGF2α activity, is pharmacologically distinct from the actions of AGN-

204396.5

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Figure 1.36 Effect of AGN-204396 (30 µM) on the contraction of feline iris produced by AL-8810.5

Because their original design is based on the structure of established TP receptor

antagonists, they possess significant off-target activity at TP receptors. However, most of

the prostamide selective tissue preparations, including the feline iris, lack expression of TP

receptors and therefore this interaction will not complicate observed pharmacology or

compromise assay results.8 Other than interactions at the TP receptor, the AGN antagonists

are quite selective for the putative prostamide receptor. AGN-204396 shows no effect on

2+ Ca signaling at 30µM on prostaglandin receptors DP, EP1, EP2, EP3, EP4, FP, and IP when activated by their agonists. EP1 actually elicits very slight agonism when treated with AGN-

204396.5

While the initial set of AGN compounds exhibit relatively decent selectivity, further extensive SAR study utilizing the feline iris assay led to the development of a second generation of compounds exhibiting even greater selectivity. AGN-211334 and AGN-211335 are the most notable, exhibiting a 100-fold higher potency and a significantly increased selectivity for the prostamide receptor over the TP receptor. AGN-211335 is a competitive antagonist (pA2 = 7.50) and when screened at other receptors it exhibited no antagonism at DP1-2, EP1-4, FP, and IP at 30 µM. It does antagonize TP receptors, however it is with less

affinity (Kb = 101 nM) than AGN-204396 (Kb = 14.9 nM)5 making AGN-211335 significantly

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more selective for the putative prostamide receptor.85 Interestingly, contraction of the feline

iris by bimatoprost is never completely blocked by AGN-211335. This could be due to off-

target activity for bimatoprost at FP receptors.

1.4.1.4 Cells that Express Prostamide-Selective Receptors Alone

Figure 1.37 The effect of (a) bimatoprost, PGF2α, 17-phenyl PGF2α, latanoprost, and latanoprost free acid and of (b) bimatoprost after pretreatment with AL-8810 (30 µM) or SQ29548 (1 µM) on peak fluorescence of Ca2+ in human T lymphoblasts (Molt-3).77

In an attempt to identify a prostamide-sensitive preparation devoid of prostanoid FP

receptors, researchers studied human Molt-3 T-cells and found that bimatoprost enhanced

2+ Ca signaling by two to three-fold over vehicle at 40-50 µM while PGF2α, 17-phenyl PGF2α,

latanoprost, and latanoprost free acid increased Ca2+ mobilization only by about one fold up

to 50 µM.77 (Figure 1.37a) While these doses are quite large, there does seem to be some

selectivity for bimatoprost over the prostaglandin F2α analogs in Molt-3 cells. The enhanced

Ca2+ signaling by bimatoprost is not significantly affected by either the FP receptor antagonist, AL-8810 (30 µM) or TP receptor antagonist, SQ-29548 (1 µM) at any concentrations up to 50 µM. (Figure 1.37b) Quantitative reverse transcription polymerase chain reaction (RT-PCR) did not detect any expression of FP receptor in human T

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lymphoblasts (Molt-3) using a probe that amplified the gene sequence present in all six FP

variants.77

Similar tests were done to determine whether bimatoprost has any actions on human

cultured osteoblasts (hTOB), where FP receptor gene was found to be expressed.

2+ Bimatoprost and AL-8810 were found to be ineffective at altering Ca signaling while PGF2α and latanoprost free acid both enhanced Ca2+ mobilization (two-three fold). As a result, this

study may represent the first evidence that bimatoprost-induced mobilization of Ca2+ in human cultured lymphoblasts (Molt-3) does not involve FP or TP receptors.

1.4.1.5 Differences in Ophthalmological Pharmacology

Finally, to emphasize the pharmacological differences between prostamide and

prostaglandin glaucoma therapeutics studies were done to prove that the mechanism of

action with which bimatoprost increases the aqueous humor outflow is unlike that of the

prostaglandins.98 Patients that were unresponsive to latanoprost therapy are successfully treated with bimatoprost suggesting pharmacological distinction between the two drugs.78

In addition, bimatoprost exhibits fewer side effects by achieving a lower hyperemic response that latanoprost and travoprost.79

1.4.2 Evidence of Prostamide Biological Actions Identical to Those of Prostaglandin

Some experts in the field of ophthalmology make the argument that bimatoprost’s intraocular pressure lowering effects are due entirely to its interactions at the FP receptor whether through direct activation by bimatoprost or through initial hydrolysis to the free acid.80 They postulate that prostamides are simply prodrugs of the prostaglandins or are

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themselves merely prostaglandin-mimetics. However it must be remembered that while this

theory may indeed be the case with anti-glaucoma prostamide agents, it does not

necessarily extend to all prostamide action throughout the body. The evidence that follows

is mostly limited to the study of prostamides only in regards to ophthalmology and the

effective treatment of glaucoma.

1.4.2.1 Isolated Receptor and Functional Tissue Studies

Interestingly, much of the same data from Woodward’s lab at Allergan Inc is presented as evidence that prostamides do in fact act on prostaglandin receptors under physiological conditions. The discrepancy comes generally in the interpretation of the results. While Woodward argues that the affinity of prostamides such as bimatoprost for prostaglandin FP receptors is too low and that an unacceptably large amount of bimatoprost would need to be used in order to sufficiently activate FP receptors,4,6,79 Sharif’s laboratory

at Alcon Research Ltd claim that the concentration of bimatoprost after topical dosing is

more than adequate to act on FP receptors.14,17,80 Relatively moderate affinities of bimatoprost were established for binding cat lung FP receptor (IC50 = 254 nM) and bovine

6 2+ corpus luteum FP receptor (Ki = 6.3 µM). In further studies, Ca mobilization was enhanced by bimatoprost in cloned human ciliary muscle FP receptor in mouse and rat cells, and phospholipase C-mediated phosphoinositide was activated in human ciliary muscle,

trabecular meshwork, mouse, and rat cells at considerably modest potencies (EC50 = 1.4-

16,88,97 3.9 µM and 0.6-9.6 µM respectively) Because reported peak levels (Cmax) of bimatoprost after topical dosing in monkey ciliary body, 0.85 µM-2.6 µM,6,79 are well within

this range of IC50 and Ki values, it is expected that bimatoprost itself would be capable of FP activation.

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1.4.2.2 Super-Sensitivity and Super-Coupling of FP Receptors in Tissue

In response to the identification of feline iris tissue being a tissue containing putative

prostamide receptors, Sharif theorizes that is it not a co-expression of a putative

prostamide receptor along with the FP receptor within tissues that preferentially recognize

prostamides.80 Instead, this exquisite sensitivity may be due to super-sensitivity and super-

coupling of the FP receptors to the signal cascade pathways in these tissues such that

bimatoprost itself, as opposed to the free acid, would activate the FP receptor more

efficiently than in other cells. In addition, amplification of the signal cascade could be

elevated and rapid so that only partial occupancy of the receptors is necessary to give full

response in the tissue. In this case even low nanomolar concentrations of bimatoprost

would effectively activate the FP receptor cascade. The establishment of FP receptor

antagonist/partial agonist, AL-8810, as a full agonist in the cat iris tissue concurs with this theory.86 This tissue is not only extra potent in response to bimatoprost, but latanoprost and cloprostenol also exhibit tremendous potencies (12 nM and 1.2 pM respectively).15 Thus it seams cat iris sphincter muscle is enriched in classical FP receptors that are extremely well coupled to their signaling cascade and do not contain a putative prostamide receptor.

1.4.2.3 FP Receptor Knockout

The most persuasive evidence to this point that prostamides do not obtain their activity through activation of receptors diverse from the prostaglandin receptors is the inability of bimatoprost to lower intraocular pressure in FP knockout mice (Figure

1.38).19,20 Therefore an intact FP receptor gene is needed in order for bimatoprost and the prostamides to elicit their pharmacological effects in vivo. Excluding the singular recent study of Molt-3 T-cells discussed previously,77 prostamide activity has not been demonstrated in the absence of FP receptor activity. A feature of all prostamide studies is

83 that PGF2a-EA effects in cells and tissues are accompanied by FP-like activity. These two

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observations give credence to the theory that prostamides like bimatoprost are indeed

acting solely on prostaglandin receptors. However, it is also very possible that this means

simply that the prostaglandin FP receptor gene encodes for the classical FP receptor and one

or several splice variants that selectively recognize the prostamide series. This alternative

splicing variant could represent the putative prostamide receptor as described by

Woodward’s lab. Regardless of whether there is selectivity among splice variants, it has thus

been established that FP receptor proteins are required for bimatoprost and its congeners to

elicit their functional biological effects.

Figure 1.38 Difference in IOP between eyes treated and untreated with bimatoprost in wild-type (solid diamonds) and FP-knockout mice (solid squares).19

1.4.2.4 Confocal Microscopy Studies

There is opposition to the conclusions drawn from the findings in both the confocal microscopy and selective antagonist studies discussed previously. In response to the confocal microscopy studies discussed previously, Sharif maintains that the multitude of PG receptors coupled to both cAMP/cGMP production and Ca2+ mobilization may mean a net

Ca2+ response is not observed.80 Likewise, the antagonism studies do not advance the

hypothesis of selectivity due to the unreasonably high concentration (30 µM) of AGN-

211334 that was used that it could interact with many different receptors. Furthermore, the

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selective prostamide antagonists might not antagonize the prostamide receptor, but instead

may inhibit conversion of bimatoprost to the free acid by FAAH.

1.4.2.5 Similarities in Ophthalmological Pharmacology

Finally, to support the position that bimatoprost and the prostamide analogs exhibit

the same pharmacology as the prostaglandin therapeutics, proponents site the similar long

term effects and side effects of these glaucoma treatments. Bimatoprost has shown the

same long term effects as latanoprost on monkeys including enlarged uveoscleral outflow

routes in ciliary muscle by degradation of extracellular matrix, sprouting nerve cells, and

exhibiting extracellular matrix-remodeling morphological changes in the trabecular

meshwork.99 Side-effects indicative of long term prostaglandin analog glaucoma

therapeutics are also shared by bimatoprost including iris pigmentation, eyelash darkening,

pigmentation of periocular skin, increased eyelash growth, and slight thinning of the central

cornea.100 All of this provides strong evidence the bimatoprost is indeed acting analogously

to prostaglandin F2a analogs in the lowering of intraocular pressure.

Latanoprost, travoprost, and bimatoprost share similar pharmacological profiles.

They are all miotin in cats and dogs but not in humans and monkeys.79 They lower

intraocular pressure in dogs, monkeys, and humans but not in cats. The singular difference

is that bimatoprost lowers intraocular pressure in rabbits, but latanoprost does not. This

phenomenon can be explained by bimatoprost acid being a potent EP1 receptor agonist

(EC50 < 3 nM). This is in discordance with Woodward’s theory that this differentiation

represents a unique action of bimatoprost resulting from interactions with a putative

prostamide receptor.

When bimatoprost and latanoprost are co-administered there is a lack of additive effects.101 This is evidence that the two drugs are competing for the same receptors. If

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bimatoprost were acting on its own receptors disparate from the FP receptors, one would

expect a synergistic effect in the lowering of intraocular pressure when both bimatoprost

and latanoprost are applied. The lack of this additive effect again emphasizes that both

these prostamides and prostaglandins are interacting at the same FP receptors.

The acute side effects associated with PGF2α analog glaucoma drugs such as

hyperemia and irritation are due to interactions at EP, DP, TP receptors present on the

ocular surface and in tissues within the eye. Side effects are not necessarily due only to

poor permeability of prostaglandin analogs through the ocular endothelium as initially

postulated. This is in accordance with the fact that ester prodrugs of PGF2α, which are more

lipophilic and would therefore pass through the ocular endothelium at a much higher rate,

are ineffective. Therefore, it is not the amide head group that gives bimatoprost its

enhanced pharmacological profile over the prostaglandins. The 17-phenyl moiety is the key

to making PGF2α sufficiently selective for FP over EP, DP, and TP receptors and thereby reducing hyperemia and irritation to an acceptable level.80

1.4.3 The Compromise: FP Receptor Splice Variants

In light of the pivotal discovery that FP receptor gene (PTGFR) knockout mice do not respond to bimatoprost and considering the prostamides’ reduced activity at prostaglandin receptors, the possibility arises that bimatoprost may interact with PTGFR products that are related but not identical to wild type FP receptors. Splice variants encoded by the FP receptor might exist and include slightly different amino acid sequences and secondary binding structures from the native FP receptor protein. Homodimerization of these protein isoforms or heterodimerization with wild type FP receptors may represent the putative prostamide receptor that has been proposed. Indeed, six alternative splice variants of the

FP prostaglandin receptor (altFPs) have been identified all containing truncated amino acid sequences with variation at the C terminus.85 Their sequences diverge at leucine 266 and

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while the wild type sequence goes on to form a third extracellular loop, a seventh

transmembrane domain, and an intracellular carboxyl terminus, the altFP receptor proteins

lack all of these features. All of their carboxyl termini are extracellular, and they contain

only six transmembrane domains. (Figures 1.39 and 1.40) pertinent

Figure 1.39 Comparison of the divergent portions of the amino acid sequences of wild type and alternative splice variant FP receptors.

leu266

FP altFP

Figure 1.40 Representation of the seven transmembrane helices of the FP receptor and the truncated six transmembrane helices with extracellular carboxyl terminus of the alternative splice variant FP receptors.

Cells transfected with each of the altFP receptors were tested against PGF2α and

bimatoprost in Ca2+ mobilization studies. The results revealed that none of the altFP receptor homodimers expressed were acted on by either 10 µM PGF2α or bimatoprost

(Figure 1.41a).85 Alternatively, cells co-transfected with wild type FP receptors and each of

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a PGF2α bimatoprost

b

c

Figure 1.41 Fluorescence traces of Ca2+ signaling in HEK cells expressing FP and/or altFP4 following

treatment with either PGF2α or bimatoprost (100 nM). (a) The effect of PGF2α (left) and bimatoprost

(right) in altFP4 expressing cells. (b) The effect of PGF2α (left) and bimatoprost (right) in FP and altFP4 85 coexpressing cells. (c)The effect of PGF2α (left) and bimatoprost (right) effects in FP expressing cells.

2+ the altFP receptors all showed a Ca mobilization response to both bimatoprost and PGF2α

(Figure 1.41b). In cells co-expressing wt FP and altFP4 receptors, this response was

observed using as little as 0.1 µM of either bimatoprost or PGF2α. Notably, cells expressing only wild type FP homodimers recognized PGF2α but not bimatoprost (Figure 1.41c). Most

of the work done with wild type and splice variant FP heterodimers following these tests

focused on the interaction and pharmacology of the altFP4 variant. The altFP4 variant

protein was selected due to its longer extracellular C terminus domain for which design of a

specific antibody might prove easier.

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An interesting observation throughout these studies involved a distinctive response

of the cells co-expressing altFP4 and wt FP to bimatoprost. Bimatoprost uniquely elicited a

second phase Ca2+ wave after the initial flux (Figure 1.41b).85 This observed effect agrees with the Ca2+ signaling studies in cat iris sphincter suggesting a native FP-altFP4 receptor

7 population exists in cat iris sphincter cells. PGF2α alternatively exhibited a steady state increase in Ca2+ signaling. The prostamide antagonist AGN-211335 selectively blocks the second phase Ca2+ mobilization of bimatoprost but not the steady-state phase of PGF2a Ca2+ signaling (Figure 1.42).85 The mechanism for this diversity in Ca2+ signaling pathways is

unknown at this point, however the data strongly correlates to the cat iris tissue studies.

2+ The divergent secondary Ca signaling pathways of bimatoprost and PGF2α may translate

into the additive effect on IOP as seen with bimatoprost and latanoprost.81

a vehicle AGN211335

b

Figure 1.42 Fluorescence traces of Ca2+ signaling in HEK cells coexpressing FP and altFP with or

without prostamide antagonist AGN211335 and following treatment with either PGF2α or bimatoprost (100 nM). (a) The effect of pretreatment with vehicle (left) or AGN211335 (1 µM) (right) on the signal

caused by PGF2α. (b) The effect of pretreatment with vehicle (left) or AGN211335 (1 µM) (right) on the signal caused by bimatoprost. Arrows indicate first the time of vehicle or AGN211335 85 administration followed by PGF2α or bimatoprost administration.

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Further study of the ciliary body confirmed these results.85 Increases in uveoscleral

outflow result from both Cyr61 remodeling of the ciliary body and phosphorylation of myocin

light chain kinase (MLC) causing ciliary muscle contraction. In wild type FP receptor

expressing cells, PGF2α incited Cyr61 mRNA upregulation and enhanced MLC phosphorylation. Bimatoprost affected neither Cyr61 mRNA nor MLC in these same cells. In cells co-expressing wild type FP and altFP4 both PGF2α and bimatoprost were able to

upregulate Cyr61 and increase MLC phosphorylation. Prostamide antagonist AGN-211335

85 blocked bimatoprost’s but not PGF2α’s effects in these cells.

While transcripts for truncated isoforms of FP receptors have been found in tissue,

this is the first example of these receptors exhibiting GPCR-like activity to ligands.102 Thus

heterodimers of the alternative spliced isoform with wild type FP receptors are possible and

uniquely recognize bimatoprost. These results suggest that the FP-altFP receptor isoforms

may be responsible for the unusual pharmacology of bimatoprost and the prostamides and

represent the putative prostamide receptor. A frequent occurrence of GPCR dimerization,

heterodimerization can create a novel ligand-recognition receptor binding site. An example

of this phenomenon is the heterodimer of prostaglandin receptors IP/TPa forming a unique

103 binding site for isoprostane E2.

Latanoprost insensitivity in some glaucoma patients has been shown to involve certain single-nucleotide polymorphisms associated with the FP receptor.104 In these

patients, bimatoprost has been proven effective.78,105 The result may represent insensitivity

of both the FP receptor and the FP-altFP receptor to PGF2α analogs, while bimatoprost

retains its activity at the alt-FP receptor. Most importantly, this alternative splicing variant

could represent the putative prostamide receptor.

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1.5 Prostamide Action on Established Proteins

Several receptors beyond the prostaglandin receptors have, in the past, been proposed as the biological targets for the prostamides. Prostamides have been screen by this and other labs at known adrenergic, cholinergic, cannabinoid, dopaminergic, prostanoid, and several other orphan receptors.79 To this point no meaningful inhibitory interactions have been found within these general screens.

1.5.1 Receptors that Recognize Anandamide

O O H N OH O N H H O HO OH

Figure 1.43 Comparison of the pharmacophoric elements of anandamide and prostamide E2. Blue circle = amide head, red squares = lipophilic chains, green triangle = conformationally restriction and polar groups

Because prostamides are the product of COX-2 metabolism of the endocannabinoid

anandamide, they contain the proper pharmacophoric elements needed for binding CB

receptors such as an amide head group and a long lipophilic tail. However, unlike anandamide they are conformationally restricted due to the introduction of the cyclopentyl moiety and contain polar hydroxyl and ketone functionality. (Figure 1.43) Therefore, while a series of PGE2 and PGF2α prostamides shown in Figure 1.44 gave enhanced binding to the cannabinoid receptors compared to the prostaglandins (0-5% inhibition at 10 µM), their potency could not rival that of anandamide (Ki = 61 nM).106 The Ki values for these

prostamides at CB1 were found to be in the range of 30 to >100 µM, while at CB2 they were found to be more potent with Ki values in the range of 1 to 10 µM.107 Because

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concentrations of prostamides are not expected to reach these levels in vivo, it can be

concluded that prostamides do not effectively bind the classical cannabinoid receptors.

Within this same study, PGE2-EA was able to stimulate G coupled proteins as

revealed by a Gs-activated increase in adenylyl cyclase. This effect was not inhibited by CB1 antagonist SR-141716A, and therefore indicates activity at some other GPCRs such as prostaglandin or other receptors. The lack of inhibition by SR-141716A also confirms a lack of prostamide action at the cannabinoid receptors.107 However, in a much more recent study of isolated human ciliary muscle tissue, the contractive effects of bimatoprost was effectively antagonized by CB1 receptor antagonists SR-141716A and, from our own lab,

AM-251. Thus the unique interactions of bimatoprost in glaucoma treatment may indeed partially be a result of interactions with the cannabinoid receptors.

HO H O H N N OH OH O O HO HO OH OH

O HO H H N N OH OH O O HO HO OH OH

HO H O H N N OH OH O O HO HO OH OH

Figure 1.44 PGF2α and PGE2 analogs tested against CB1 and CB2

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Prostamides were considered possible ligands for vanilloid receptors, again due to

their derivation from the vanilloid receptor agonist anandamide. Of the prostamides E2, D2,

2+ and F2α, only prostamide F2α was able to weakly act on TRPV1 in a Ca mobilization assay.

2+ With an EC50 of 15.0 µM, it had a maximal effect 31% of that of 4 µM ionomycin, a Ca

8 mobilizer. In a separate study, PGE2-EA at 10 µM gave only 23.58% displacement of specifically bound [3H]-resiniferatoxin from TRPV1.9 In light of this poor affinity of the prostamides for the vanilloid receptor, it is not reasonable to consider them agonists at relevant concentrations.

1.5.2 Peroxisome Proliferation-Activated Receptor

Some pertinent findings have led to the possibility that prostamides may activate the peroxisome proliferation-activated receptor γ (PPARγ).108 Anandamide has long been known

as an immunomodulator and controller of cytokine production, however more recent studies

have shown that this effect is not exclusively a result of interaction at cannabinoid

receptors.108,109 Anandamide inhibits interleukin-2 (IL-2) secretion in a concentration-

108 dependent manner (IC50 = 11.4 µM). (Figure 1.45a) Interleukin-2 is an

autocrine/paracrine factor secreted by activated T-cells and promotes T-cell proliferation,

and inhibition of IL-2 signifies immunosuppression.110 IL-2 inhibition by anandamide could

not be reversed by addition of a combination of CB1 inhibitor SR-141716A and CB2 inhibitor

SR-144528, and thus anandamide effects on IL-2 are not mediated by the cannabinoid

receptors.108 (Figure 1.45b) Alternatively, inhibition of IL-2 was partially reversed by

pretreatment with COX inhibitors flurbiprofen and piroxicam and COX-2 selective inhibitor

NS-398. (Figure 1.45c,d) Therefore metabolic conversion of AEA to the prostamides by

COX enzymes is necessary for the inhibition of IL-2, and at least some of anandamide’s

known immunosuppressive effects may be attributed to its prostamide metabolite.

Pretreatment with PPARγ-

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a

c b

e d

Figure 1.45 Effect on IL-2 levels in PMA/ionomycin-stimulated murine primary splenocytes by (a) increasing concentration of anandamide, (b) increasing concentration of CB1 and CB2 inhibitors SR141716A and SR144528 in the presence and absence of anandamide, (c) increasing concentration of anandamide in the presence and absence of COX inhibitor FBN, (d) increasing concentration of anandamide in the presence and absence of COX-2 selective inhibitor NS398, and (e) increasing concentration of PPARγ selective inhibitor T0070907 in the presence and absence of anandamide.108

specific antagonist T-0070907 also partially antagonized the suppression of IL-2 secretion exhibited by AEA, leading to the conclusion that prostamide is a ligand for the PPARγ.

(Figure 1.45e) This is a reasonable assertion as the generally accepted, naturally occurring

111 activating ligand for PPARγ is 15-deoxy-Δ-prostaglandin J2. This study asserts that

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prostamides are the endogenous compounds responsible for at least some of the

immunomodulatory effects observed with anandamide administration and that these effects

including inhibition of IL-2 are the result of binding the PPARγ nuclear receptor.108

1.5.3 Lipid Metabolizing Enzymes

The possibility has been proposed that prostamides achieve their biological

significance through interactions at key lipid metabolizing enzymes. Prostamides themselves

may inhibit certain enzymes thereby increasing local levels of certain lipid mediators and

decreasing levels of their metabolites. Similar to the receptor studies, the enzymes that

would logically be appropriate for interactions with prostamides are those that act on their

endocannabinoid precursor anandamide.

FAAH is the main protease responsible for the degradative metabolism of

anandamide. If prostamides were to act as substrates for FAAH, they would inhibit the

inactivation of anandamide and thus increase endocannabinoid tone. In this way,

prostamides biological purpose would in effect be to regulate cannabinoid activity. If FAAH

were to act on the prostamides as substrates, the metabolite produced would likely be the

correlating prostaglandin. In a similar fashion, cellular uptake of the endocannabinoids by a

transporter protein is the first step in anandamide inactivation and inhibition by the

prostamides would lead to higher local concentration of anandamide.8

Tissue specific hydrolysis by FAAH may explain why prostamides have similar

potency to prostaglandins in certain specific assay preparations such as the cat iris79 and

guinea pig.9 Assuming the coexistence of prostaglandin and prostamide receptors, tissues expressing lower levels of FAAH would mean an increase in prostamides available to elicit the contractile effects within these tissues. Conversely, assuming that there is no putative prostamide selective receptor, an upregulation of FAAH would equate to a greater degree of

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metabolism to free and active prostaglandins. Thus this hypothesis does nothing to prove or

disprove the existence of prostamide receptors in these tissues, but only concerns the effect

of FAAH regulation on tissue responses.

All of these suggestions are void however in the case of the prostamides as they are

not acted on by FAAH and therefore do not inhibit the hydrolysis of anandamide.79 None of

the prostamides E2, D2, or F2α significantly inhibited FAAH’s activity in the hydrolysis of

[14C]AEA. (Figure 1.23a) They also did not inhibit the uptake of AEA by RBL-2H3 cells in a substantial way and do not undergo cellular uptake by rat brain synaptosomes or intact

RBL-2H3 cells.8 (Figure 1.46) In a similar study, the level of inhibition of specific binding of

100 nM anandamide (32.54%) to CB1 was not enhanced by PGE2-EA at concentrations up to

9 10 µM. (Figure 1.23b) Therefore PGE2-EA is not inhibiting FAAH and in so doing does not

increase cannabinoid tone.

Figure 1.46 Lack of inhibitory effect of prostamides E2, F2α, D2 and bimatoprost (50 µM) on cellular uptake of [14C]anandamide by intact RBL-2H3 cells8

The possibility exists that prostamides could simply be the product of an inhibitory

interaction by its precursor anandamide. Because anandamide is a substrate for COX-2, it is

quite possible that anandamide acts to regulate prostaglandin production and prostamides

are merely the products of this interaction. Anandamide would act as an inhibitor of COX-2

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112 biosynthetic pathway. Likewise, prostamides and their COX-2 intermediate PGH2-EA are substrate/inhibitors for the individual prostaglandin synthases thereby regulating production of the prostaglandins. Thus, there is a well-established and widely accepted competition between prostamide biosynthesis and prostaglandin synthesis. However, the likelihood that this is the only justification for prostamide’s presence within the body does not seem probable.

Another interesting aspect to consider is the suggestion that the inhibition of the metabolic enzymes for the biosynthesis and degradation of prostamides may mean that prostamides play a substantial role in the therapeutic actions of several drugs. Selective

COX-2 inhibitors may elicit some of their effects by reduction of not only prostaglandin levels but also prostamide levels. Accordingly, inhibition of prostamides may greatly contribute to the observed beneficial and toxic pharmacology of the coxibs discussed in

Section 1.2.3.

FAAH inhibitors such as PMSF, URB-597, and PF-622 have been used mainly to increase anandamide concentrations and, in turn, increase activation of cannabinoid receptors. However, FAAH inhibition in this way will also lead to a significant increase in prostamides due to the greater availability of anandamide to be acted on by COX metabolism. Therefore, when studying the pharmacological actions of FAAH inhibitors, increased prostamide production must be taken into consideration in addition to the given endocannabinoid interactions.

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1.6 Prostamides in Inflammation and Other Systems

1.6.1 Presence of Prostamides at Physiologically Relevant Concentrations In Vivo

It is uncertain whether prostamides exist within the body at concentrations that would make them relevant for pharmacological interactions. To this point, no study has proven the endogenous presence of any prostamide at basal levels. However, it is likely that under certain physiologic conditions such as during inflammatory or infectious challenge that anandamide release and COX-2 up-regulation could occur to synergistically produce a relevant amount of prostamides.

Prostamides are quite stable especially when compared to their prostaglandin counterparts. While prostaglandins have substantial biological effects, they exhibit very short half-lives in vivo and therefore only function as paracrine or autocrine messengers.113

The initial step in metabolic deactivation of the prostaglandins is oxidation of the 15- hydroxyl group by 15-hydroprostaglandin dehydrogenase. Because the prostamides are not efficient substrates for this enzyme, the prostamides have a much longer half-life than the prostaglandins. After a 2 mg/kg intravenous dose in rats, PGE2-EA was detectable up to 2

hours after dose.3 Exhibiting large volume distribution and a half-life of over 6min means

that prostamide may be sufficiently stable to exert endocrine activity. This extra metabolic

stability may mean that prostamides serve as latent sources of prostaglandins at sites

remote from their tissue of origin.

The challenge in maintaining a relevant concentration of prostamide is not dependent

on the stability of prostamides or the effective metabolism by COX-2, which is as efficient as

arachidonic acid metabolism,1,2 but instead on the stability of the precursor, anandamide.

FAAH hydrolyzes anandamide to arachidonic acid so effectively that there may not be

sufficient concentrations present to allow for biosynthesis of prostamide. In addition to the

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fast metabolism of anandamide, the prostamide biosynthetic pathway also suffers due to

the fact that the endogenous pool of anandamide is substantially less than arachidonic acid.

In human plasma, anandamide exists in the low nanomolar range.109 Due to the lack of anandamide presence in the lipid layer, production of a sufficient level of prostamide for detection is more challenging.

Prostamide production in vivo has been established via HPLC-MS following an external intravenous dose of 50 mg/kg anandamide in mice.114 This proves that there exists

a pathway within the body to convert anandamide to the prostamides. While the resulting

concentration of total prostamide (F2α, E2, D2) 30min after dosing was low 0.36 – 3.17 ng/g, when tested in FAAH knockout mice there was a much larger concentration 0.9-25.9 ng/g.

This result is not due to a lack of FAAH hydrolysis of the prostamides but instead due to the increased amount of precursor anandamide.

1.6.2 Prostamides in Inflammation, Neuroinflammation and Neuroplasticity

Several studies have shown that under inflammatory or infectious challenge COX-2

expression is up-regulated and anandamide release is stimulated.55-58,115,116

Lipopolysaccharide and interleukin-1β have both been shown to incite increases in anandamide and COX-2,56-58 and because a resulting increase in prostamide synthesis is expected under these conditions, prostamides may play an important role as lipid mediators in inflammation and infection. In the subsequent Section 1.6.3, it will be shown that WISH cells and primary amnion tissue are able to produce prostamide E2 after treatment solely with anandamide, however the production is greatly enhanced by the presence of COX-2 induction by IL-1β.55 When RAW 264.7 macrophages were treated with lipopolysaccharide a

117 significant production of PGE2-EA was incited. In fact, one study found that anandamide

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stimulation itself gives substantial induction of COX-2.115 Thus regulation of prostamides

could entail therapeutic indications in inflammatory disease states such as rheumatoid

arthritis, asthma, periodontitis, psoriasis, irritable bowel syndrome, Crohn’s, coeliac, and

Alzheimer’s diseases, and even cancer.

Several recent studies provide evidence that prostamides are neuroprotective and

may have an important role in neuroplasticity and learning.109,118,119 Neurodegenerative

processes are often accompanied by inflammation, and therefore the theoretical increase in

prostamide formation resulting from an increase in anandamide production and COX-2

expression could represent a mechanism of neuronal survival in damaged nerve tissue.

One such study monitored the neuroprotective and antitumor activity of AEA and

118 + PGE2-EA. Using K reduction within the cell nutrient medium to trigger apoptosis of

neurons, the effects of treatment with either AEA or PGE2-EA on cell survival was measured.

While AEA treatment resulted in no neuroprotection up to 1 µM, PGE2-EA show a potent

increase in neuronal survival. The prostamide exhibited a 21% increase in survival at 0.1

µM, 59% increase at 0.5 µM, and 77% increase at 1.0 µM.

The effects of AEA and PGE2-EA on tumor cell apoptosis were also observed in this study. Rat C6 glioma cells were treated with various levels of AEA and PGE2-EA, and it was

found that while AEA showed substantial antitumor activity (20% reduction at 2.5 µM, 25%

at 5 µM, 35% at 10 µM), PGE2-EA showed no signs of attack on tumor cells up to 5 µM.

Therefore, the metabolism of AEA to PGE2-EA by the increased COX-2 expression exhibited within some primary tumors and metastases serves as a pathway of inactivation of AEA’s antitumor effects. PGE2-EA does not distinguish in its protection of neuronal cells as it both

protects normal cells and does not destroy tumor cells.

Prostamide E2 has recently been found to have an essential role in the regulation of interleukin-12 (IL-12), a heterodimeric cytokine that regulates innate immunity and is

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produced mainly by monocytes, macrophages, dendritic cells, and brain microglia.109 IL-12

has been shown to play a crucial role in the initiation and perpetuation of various

autoimmune and chronic inflammatory demyelination diseases such as multiple sclerosis

(MS).120-122

During neuroinflammation endocannabinoids are released, mainly by microglial cells,

and exert neuroprotective actions. In order to determine whether some of this

neuroprotection is due to anandamide interactions with IL-12 or related protein IL-23,

researchers looked at both anandamide and its COX-2 metabolite prostamide E2 in activated macrophages and microglial cells.109 mRNA expression of p19, p35, and p40 subunits of IL-

12 and IL-23 was monitored upon treatment of lipopolysaccharide (LPS) and interferon- gamma (IFNγ)-activated microglial cells. (Figure 1.47)

Figure 1.47 (a) Administration of AEA (10 µM) inhibits the mRNA expression of p35, p40, and p19 in microglial cells activated by LPS/IFNγ (b) ELISA analysis shows suppression of IL-12p40 expression by microglial cells is dose-dependent on AEA administration.109

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Anandamide is able to reverse the induction of p40, p35, and p19 by suppressing IL-

12p40 promoter gene activity. These effects occur independently of activation of CB1, CB2,

109 and TRPV1 and thus are not affected by the respective antagonists of these receptors.

Importantly, prostamide E2 (10 µM) is equally effective reversing the LPS and IFNγ-induced

upregulation of p40, p35, and p19 mRNA expression. (Figure 1.48) It decreased the p40

promoter gene activity in a dose-dependent manner. Both anandamide and prostamide E2 action is facilitated through activation of the EP2 receptor. The EP2 antagonist, AH-6809, was used to partially reverse the inhibitory effects of both AEA and PGE2-EA on transcriptional

activity of IL-12p40 in RAW 264.7 cells. EP4 is not involved, as EP4 receptor antagonist AH-

23848B had no reversal effect. These results were confirmed on the protein expression level

as AEA and PGE2-EA both down-regulated the induction of IL-12p40 protein by LPS and

IFNγ-activated microglial cells.

a b c

Figure 1.48 (a) Administration of prostamide E2 (10 µM) inhibits the mRNA expression of p35, p40, and p19 in microglial cells activated by LPS/IFNγ (b) Suppression of IL-12p40 expression by RAW 264.7 cells is dose-dependent on prostamide E2 administration. (c) Administration of EP2 selective inhibitor AH6809 reverses the effects of both AEA and prostamide E2 on IL-12p40 suppression in RAW 264.7 cells.109

Remarkably, inhibition of COX-2 by selective COX-2 inhibitor, NS-398 (10 µM),

reverses the effects of AEA on both p40 promoter activity and IL-12p40 expression,

signifying that anandamide’s regulation of IL-12 is exclusively the result of its COX-2

metabolism to PGE2-EA. Use of NS-398 along with PGE2-EA did not change its effective

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inhibition of p40 promoter activity. This represents the mechanism by which AEA has been

shown to reduce the development of experimental models of MS including chronic relapsing-

remitting experimental autoimmune encephalomyelitis (CREAE) and Theiler’s murine

123-125 encephalomyelitis virus (TMEV). Within this context, prostamide E2 is acting exclusively as a prostaglandin mimetic and it has very recently been established that

126 prostaglandin E2 has similar effects on IL-12 regulation.

Prostamide E2, like prostaglandin E2, regulates neuroplasticity and learning by dynamically maintaining membrane excitability, synaptic transmission, integration, and plasticity in the hippocampus. In opposition to the effects of anandamide, prostamide E2 enhances hippocampal long term potentiation (LTP).119 Whether all of these effects are the

result of prostamide interactions at prostaglandin receptors alone or at other unknown

receptors has yet to be determined.

1.6.3 Misidentification of Prostamides as Prostaglandins

Figure 1.49 Percentage of total compound identified to be “PGE2” by RIA that is actually PGE2 (open

bars) and that is PGE2-EA (solid bars) as a result of treatment of WISH cells and amnion with IL1 and anandamide55

Complicating the study of prostamides is the lack of selective methods of

identification used in many studies of the prostaglandins and prostamides. There is

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significant concern that studies using techniques such as radioimmunoassay to measure the

presence of prostaglandin in a system may instead give the additive concentration of both

prostaglandins and prostamides together. This gives rise to the risk that some

pharmacological properties previously attributed to prostaglandins may indeed be the result

of prostamide production as well or instead. One study gave evidence of this possible

misidentification and demonstrated that using a radioimmunoassay (RIA) alone to detect

and quantify prostaglandins is not sufficient to differentiate between prostaglandins and

55 prostamides. Both PGE2 and PGF2α antibodies show major cross reactivity with the prostamide counterparts. In fact, not only will these antibodies recognize PGE2-EA, but all

antibodies tested showed a significantly higher affinity for PGE2-EA than for PGE2. Affinity

for PGF2α-EA and PGF2α were very similar. (Figure 1.49) When compound identified by standard RIA to be purely PGE2 underwent thin layer chromatography (TLC), a substantial

amount of and PGE2-EA was isolated. (Table 1.5)

Table 1.5 Percentage of PGE2 in RIA determined to be PGE2-EA following purification by TLC Found for WISH cells

Basal levels 2%

Upon treatment with 3% 0.2ng/mL IL-1β

Upon treatment with 95% 0.2ng/mL IL-1β and 10µM AEA

Upon treatment with 77% 10µM AEA

Found for amnion explants

Basal levels 24%

Upon treatment with 42% 10µM AEA

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The study concludes that chromatographic or mass spectrometric methods in addition to RIA are necessary for detecting and quantifying both prostaglandins and prostamides independently of each other. It also raises the possibility that studies up to this point have mistaken pharmacological effects of the prostamides and attributed them to prostaglandins.55

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123. Baker, D.; Pryce, G.; Croxford, J. L.; Brown, P.; Pertwee, R. G.; Huffman, J. W.; Layward, L., Cannabinoids control spasticity and tremor in a multiple sclerosis model. Nature 2000, 404, 84.

124. Arevalo-Martin, A.; Vela, J. M.; Molina-Holgado, E.; Borrell, J.; Guaza, C., Therapeutic action of cannabinoids in a murine model of multiple sclerosis. J. Neurosci. 2003, 23, 2511.

125. Croxford, J. L., Therapeutic potential of cannabinoids in CNS disease. CNS Drugs 2003, 17, 179.

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126. Van Elssen, C. H. M. J.; Vanderlocht, J.; Oth, T.; Senden-Gijsbers, B. L. M. G.; Germeraad, W. T. V.; Bos, G. M. J., Inflammation restraining effects of prostaglandin E2 on natural killer-dendritic cell (NK-DC) interaction are imprinted during DC maturation. Blood 2011, 118, 2473.

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Chapter 2: Design and Synthesis of α-Head and ω-Tail Modified Prostamide E2 Analogs

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2.1 Analog Design and Rationale

The necessity has been established for further elucidation of prostamides’ biological actions, independent from those of the prostaglandins. To address the scarcity of valuable prostamide derivatives for use in receptor screening, identification, and isolation of targeted protein interactions, a series of prostamide E2 analogs were designed. (Figure 2.1) A multiple step synthesis, presented in Section 2.2, was devised that allows for convergent functionalization of both the head and/or tail groups providing these potentially useful probes for characterization studies. Several factors were taken into consideration during the selection of analogs to be synthesized.

Figure 2.1 Prostamide analogs selected for synthesis

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2.1.1 Configuration about the Cyclopentyl Ring

First among these considerations were changes in the configuration of the

prostamide central cyclopentyl ring. While cyclooxygenase metabolism of anandamide is

able to produce all of the standard configurations corresponding to the prostaglandins

including PGD2-EA, PGE2-EA, PGF2α-EA, PGI2-EA, the major product of prostamide

1,2 biosynthesis, by far, is PGE2-EA. (Figure 2.2) Thus, all of the analogs chosen for

synthesis share the E2 configuration about the cyclopentyl moiety.

X H N OH O Y OH O OH OH O OH O N H R1 R1 R1

O R2 HO R2 HO R2 HO R2 - - α- PGD2 EA PGE2 EA PGF2 EA PGI2-EA

Figure 2.2 Known prostamide products of cyclooxygenase metabolism of anandamide. Prostamide E2 (PGE2-EA) is the major product

2.1.2 Covalent ω-Tail Modifications as Affinity Probes

Secondly, this set of compounds incorporates azido and isothiocyanato functionalities

at the terminal tail position as modifications for covalent binding to the putative prostamide

receptor. These covalent probes, also known as affinity labels, are commonly used in the

characterization of enzyme, receptor, and transporter proteins.3-6

2.1.2.1 The Photoactivatable Azido and the Electrophilic Isothiocyanato Groups

The azido functionality falls into the category of photoactivatable ligands which are chemically inert until photoirradiation converts them into reactive nitrene or carbene

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intermediates.7,8 (Figure 2.3) The reactive intermediate can then form a covalent bond with nearby amino acid residues.9-12,13,14 Other photoaffinity ligands, which can take the

form of carbenes when photoirradiated, include diazirines and α-ketodiazo groups.15,16 By preequilibrating the photoaffinity probe with a membrane thought to express prostamide receptors and subjecting it to shortwave UV irradiation (λ = 287 nm, 216 nm; ε = 25 M-1cm-

1),17 an irreversible cross-link is established to the membrane proteins that recognize the

prostamide probe. The result is one or more ligand-protein adducts, of which one or more may be prostamide-selective receptors.

Figure 2.3 Mechanism of covalent attachment by the azido photoaffinity label

Figure 2.4 Mechanism of covalent attachment by the electrophilic isothiocyanato group

Similarly, the electrophilic isothiocyanato group is capable of covalently labeling the binding site of a target protein. Isothiocyanate is a moderately reactive electrophile, inert in water, but prone to reaction with electron-rich residues such as those bearing sulfhydryl,

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unprotonated amino, imidazolyl, or hydroxyl under physiological conditions.18 (Figure 2.4)

Thus, cysteines, lysines, histidines, or serines in the vicinity of the receptor active site may be capable of nucleophilic reaction with the isothiocyanato group. The nucleophilicity of such residue side chains is most likely enhanced in a catalytic fashion by interactions with

proximal amino acids. In the case of cysteine, the complex formed is expected to be the

thiocarbamate. Other electrophilic probes capable of binding covalently with target proteins

include Michael acceptors, haloacetamides, aldol esters, and nitrogen mustards.19

Electrophilic probes have been employed extensively as tools for the study and characterization of a series of receptors including the opioid,20,21 NMDA,22 sigma,23 and

benzodiazepine24 receptors.

2.1.2.2 Utility of Covalent Affinity Probes

By covalently modifying the proteins to which they bind, affinity labels can aid in the detection and isolation of prostamide binding sites within a given membrane preparation.

Following incubation and activation of the probes bound to the membrane proteins and

washing to clear unbound compound, specific binding assays can be run using tritiated

3 prostamide ligand ([ H]PGE2-EA). If ligand binding is significantly inhibited, the affinity probes are indeed covalently binding one or more proteins within the membrane. To isolate

these proteins, bifunctional probes containing the isothiocyanato or azido covalent

modifications and a radiolabel attachment such as 125I can be employed. Using this

bifunctional radiolabelled affinity probe essentially allows for tagging the target proteins

with radiolabel that can then be used to aid in purification of the prostamide-binding

proteins from other proteins within the same membrane preparation.

Additionally, affinity labels provide useful information about the receptor distribution, molecular weights, and amino acid residues in the vicinity of the active site. This further

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characterization of the receptor, including verification of the identity of the labeled amino acid residue bound to the probe, is revealed through purification of the covalently linked ligand-receptor complex, followed by digestion of the entire protein, and analysis of the individual peptide fragment using sequencing or mass spectrometry. These studies aim to identify and characterize the binding domain within the receptor by gaining direct information about the structural features of the receptor binding site. Information gained would assist in the design of improved receptor probes and ultimately potent and selective prostamide receptor agonists and antagonists. This methodology has been used for studying a number of receptors including NMDA,25 muscarinic,26 a1-adrenergic,27 d2-dopamine,28 serotonin,29 and retinal30 receptors.

The first obstacle in this process of receptor protein detection and isolation is the

design of probes with sufficiently high affinity for binding. Before their ability to covalently

modify the binding site can be tested, potent probes must be synthesized and tested for their binding affinity. This is most essential for the photoaffinity probes which will make more effective covalent labels when their potency is high and they occupy more receptor binding sites as they undergo photoirradiation. If not fully bound, the nitrene generated may bind elsewhere on the protein or may undergo intramolecular rearrangement to the imine.31 Elevated affinity also ensures less offsite activity with other proteins at a given concentration of affinity probe. Thus, design of highly potent probes to be evaluated by SAR binding studies is the primary initiative and, once established, these probes can be used to covalently bind their target receptors to aid in isolation, identification, and characterization.

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2.1.3 α-Head Group Modifications

Figure 2.5 Head group variations included in this set of analogs. The first three have been shown to improve stability against enzymatic hydrolysis of anandamide, and the fourth is the product of cyclooxygenase metabolism of 2-Arachidonoyl glycerol (2-AG).

The next area in which alterations were made is the α head group position. (Figure

2.5) Variations in the functionality of the ethanolamide moiety have been shown by this lab

32 to increase stability of fatty acids toward hydrolyzing enzymes. Hydrolysis of anandamide

by FAAH to arachidonic acid is greatly reduced by the introduction of R- and S-2-methyl

ethanolamides and cyclopropylamide substituents in place of the ethanolamide. Additionally,

introduction of these substituents do not affect the ligand’s ability to bind the cannabinoid

receptor, and in some cases even increased its affinity. Thus, inclusion of these same

groups within the prostamide structure may similarly enhance their stability and decrease

the likelihood that their biological actions are the result of hydrolysis to prostaglandin prior

to binding.

A 2-glycerol ester head group was also incorporated into this set of compounds to

ascertain whether it is capable of exhibiting its own distinct biological actions. Similar to the

prostamides, prostaglandin glycerol esters are the biosynthetic product of metabolism by

33 COX-2 on a different endocannabinoid, 2-Arachidonoylglycerol. (Figure 2.6) With a

similarly unknown pharmacology, the prostaglandin glycerol ester makes a valuable addition to the test compounds in achieving a more thorough investigation of the COX-2 metabolic products of the endocannabinoids.

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Figure 2.6 Representation of the eicosanoids prostaglandin-glycerol ester (PG-G), prostaglandin (PG), and prostamide (PG-EA) biosynthesis from the fatty acids 2-AG, AA, and AEA respectively and the proposed action and degradation of PG-G and PG-EA.

2.1.4 The 15-Hydroxyl Stereocenter

Figure 2.7 Variation of the 15-hydroxyl stereochemistry incorporated into the proposed analogs

The stereochemistry of the hydroxyl group in the 15 position was the final analog variation. (Figure 2.7) As discussed in the following section, synthetic challenges arose in

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the stereoselective reduction of the enone precursor to give solely the S-allylic alcohol. With

all chiral methods attempted giving some proportion of R and S-allylic alcohols, both were

isolated and carried through the synthesis. Additionally, while the 15S isomer is the native

prostanoid configuration, the 15R isomers mimic the stereochemistry of the aspirin- triggered lipoxins (ATL).

Shown to have a leading role in mediating inflammation resolution, lipoxins are the biosynthetic products of metabolic oxygenation of arachidonic acid by the lipoxygenase

enzyme (LOX).34-36 Thus, the lipoxins are similar to prostaglandins and prostamides in that

they share the same precursor, arachidonic acid, and their biosynthetic pathway requires

enzymatic oxidation. The location and extent to which the oxidation occurs is entirely

determined by the identity of the enzyme which metabolizes the arachidonic acid.

Oxygenation by COX adds two molecules of oxygen, one across the 9- and 11- positions and

37,38 one at the 15- position giving PGG2, the intermediate precursor to the prostaglandins.

(Figure 2.8) Conversely, formation of the lipoxins requires addition of two molecules of oxygen by distinct lipoxygenases.39 (Figure 2.9a) Addition of oxygen across the 5- and 6-

Figure 2.8 Oxygenation of arachidonic acid by COX

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positions by 5-lipoxygenase followed by addition of oxygen at the 15-position by 15- lipoxygenase generates lipoxin A4 (LXA4). Alternatively, oxygenation across the 14- and 15-

positions by 15-lipoxygenase and addition of oxygen at the 5-position is also possible and

leads to the formation of lipoxin B4 (LXB4).

Figure 2.9 (a) Oxygenation of arachidonic acid by the lipoxygenases (LOX) to form the lipoxins LXA4 and LXB4 (b) Oxygenation of arachidonic acid by aspirin-acetylated cyclooxygenase-2 (COX2) to form the 15 epi-lipoxins also known as the aspirin-triggered lipoxins (ATLs).

Remarkably, COX2 also exhibits the ability to form lipoxins when acetylated by the

NSAID, aspirin.40 Upon treatment with aspirin, COX2’s active site is acetylated, inhibiting its

ability to produce the prostaglandins. However, its function is not entirely terminated as it is

able to oxidize anandamide to 15-epi lipoxins. (Figure 2.9b) Also known as the “aspirin-

triggered lipoxins,” these eicosanoids not only exhibit anti-inflammatory and pro-resolving

properties, but their 15R configuration makes them more potent and longer acting than the

native 15S lipoxin compounds.41 Thus, the 15R lipoxins are more stable towards metabolism and, for this reason, the R configuration of the prostamides was included in the current set of analogs.

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2.2 Prostamide Analog Synthesis

2.2.1 Challenges in Prostaglandin and Prostamide Synthesis

The total synthesis of prostaglandins and prostaglandin analogs such as the

prostamides presents numerous challenges. Establishing the absolute stereochemistry of

four (as in PGD2, PGE2) or five (as in PGF2α) asymmetric carbons is just one of the major

concerns that needs to be overcome when taking on prostaglandin synthesis. (Figure 2.10)

Additionally, five-membered rings do not tend to adopt predictable conformations, so

specific methods to introduce the chiral functionality surrounding the cyclopentyl ring will

need to be employed. Fortunately, commercially available precursors exist that already

contain the required stereochemistry and alleviate these initial dilemmas. Introduction of

the olefins in the head and tail chains require stereospecific introduction of one Z- (cis) and

one E- (trans) double bond. Using specific Horner-Wadsworth-Emmons or Wittig conditions

O α HO α 9 Z 9 R Z (R) ( ) (S) ( ) ( ) 8 COOH 8 COOH 5 1 5 1 12 20 12 20 S (S) R ( ) ω ( ) (R) 11 (R) E ω 11 13 (E) 15 13 ( ) 15 HO O OH OH PGD PGE2 2 HO α 9 R Z (S) ( ) ( ) 8 COOH 5 1 12 20 (S) (R) (R) ω 11 13 (E) 15 HO OH PGF2a

Figure 2.10 Structures of the standard endogenous prostaglandins PGE2, PGD2, and PGF2α with atoms and stereochemistry labeled.

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will allow for easily controlled institution of these alkenes. Answers for these issues have

been well established and should not prove menacing to the total synthesis being

undertaken. However, there were still some substantial obstacles faced during our

prostaglandin analog synthesis.

The most prominent challenge when dealing with prostamide synthesis, especially for

E2 analogs, is the instability of the cyclopentyl ring hydroxy groups. Throughout the

synthesis of PGD2 or PGE2 analogs, the β-hydroxyketo moiety is unstable and will readily undergo dehydration to the α,β-unsaturated ketone under either acidic or basic conditions.

Prolonged exposure of these compounds to base gives the more thermodynamically stable di-substituted α,β-unsaturated ketone through migration of the double bond. (Figure 2.11)

A careful protection scheme of the β-hydroxy functionality will need to be employed to ensure this dehydration is less likely to occur during the synthesis. Secondly, establishment of the 15-hydroxyl moiety can pose a very challenging dilemma especially due to its considerable distance from the other stereocenters. In this remote location, the preexisting asymmetry within the ring cannot be used to establish a new chiral center at the 15- position. Unique strategies for introduction of the 15S-hydroxyl group will need to be employed in order to gain optical resolution. Additionally, determination of the identities of the resulting chiral isomers will need to be addressed to assess the effectiveness of the chiral reductions.

O O O R' OH-/H+ R' OH- R'

R'' R'' R'' RO Figure 2.11 Acidic or basic dehydration of the β-hydroxyketone to the α,β-unsaturated ketone and d - the thermodynamic hydrogen shift to the stable di-substituted α,β-unsaturated ketone. (k 1OH = 16.0 -1 -1 d + -1 -1 d - -1 -1 42 M min , k 1H = 0.0043 M min ; k 2OH = 1.6 M min )

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There have been several approaches to prostaglandin synthesis throughout the years involving formation of the cyclopentane ring from acyclic precursors or from precursors containing a preformed cyclopentane nucleus.43-49 However, Corey’s bicycloheptane approach, first developed in 1969, has proven to be the most elegant and practical.50-54

With numerous improvements since its disclosure, the Corey strategy for prostaglandin synthesis allows for the synthesis of all the known prostaglandins from a common optically

active precursor. The Corey synthesis is so ubiquitous to prostaglandin synthesis that a key

intermediate comprising the cyclopentyl ring with established stereochemistry has been

named Corey’s Lactone and is commercially available from several sources. (Figure 2.12)

As with a majority of current day prostaglandin syntheses, our synthesis utilizes Corey’s

Lactone as its essential starting material. Because this compound is sold in large scale and

contains within it four of the five chiral centers required, we need only concern ourselves

with the stereoselective synthesis of the remaining hydroxyl at the 15-position.

O O O O

O O OMe O O O

Corey's Lactone Corey's Lactone Benzoate Aldehyde

Figure 2.12 Corey’s lactone isolated in the original synthesis and Corey’s lactone benzoate aldehyde the commercially available starting material for our synthesis.

2.2.2 Novelty and Advantage of the Current Synthesis

While we begin our synthesis with Corey’s established starting material and

incorporate some of his synthetic methods into our overall strategy, there is significant

novelty within the synthesis designed by our lab. The innovation lies mainly within the

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omega tail functionality and the ability to make facile modifications of this group to

numerous other moieties in one or two additional reactions. Standard prostaglandin analog

synthesis establishes the entirety of the tail chain in the very first Horner-Wadsworth-

Emmons reaction and its core and tail structure remains unchanged throughout. No current

literature demonstrates the establishment of the core prostaglandin structure, including

both long chains, followed by functionalization of the head and tail positions alone to the

give prostamide analogs proposed in this study. Additionally, using the synthetic route

established here, the tail hydroxyl moiety can serve as an entrance to a vast number of

alterations in functionality. (Figure 2.13)

Figure 2.13 Synthetic directions allowed by the key intermediate alcohol 20. (LG = Leaving group, Ox = Oxidant)

For example, the hydroxyl can act as a nucleophile in a reaction with an isocyanate or isothiocyanate to form a carbamate or a thiocarbamate. A Williamson ether reaction,

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using a strong base and introducing an alkyl halide, gives way to numerous alkyl ethers.

The alcohol can be converted to a mesylate, tosylate or triflate leaving group to be acted on by a selective nucleophile in an SN2 reaction. In addition, these leaving groups can undergo

a basic elimination to give the alkene product. This alkene can then be functionalized to

several different groups by adding across the double bond including, but not limited to,

ethers by treatment with alcohols and a cyano by treatment with HCN. Oxidation of the tail

hydroxyl to an aldehyde allows for a subsequent Wittig-type reaction or an aldol reaction to

give a product of further alkylation. Oxidation to a carboxylic acid gives entry to numerous

esterifications or amidations. Additionally, the hydroxyl can be modified directly to a

different functionality such as azido or isothiocyanato groups as we chose to do. Adopting

this novel synthesis allows for nearly endless functionalization of the alpha head and omega

tail in parallel. Thus introduction of the tail hydroxyl sets our synthesis apart from the

current established prostaglandin synthesis due to the vast number of tail analogs that are

now easily attainable.

Additionally, introducing an ω-tail functionality adds a certain level of complexity to

the typical prostaglandin synthesis that has not been attempted by any other lab to date.

The key to success in this realm is thoughtful selection of the protective groups that will

allow for deprotection of the omega tail hydroxyl in the presence of the other protected

hydroxyls. In order to ensure that selective modification of this group is attainable, careful

implementation of an appropriate protective scheme must be employed. Silyl protecting

groups are ideal for prostamide synthesis owing to their general stability under basic

conditions and to the fact that their reactivity during addition and removal is based on the

choice of silyl group substituents.55,56 The steric demand of the silyl group is directly

correlated to its protective stability as silyl ethers containing larger substituents and those

protecting higher substituted hydroxyls are more stable towards acid hydrolysis. Stability of

the common silyl ethers toward acid increases in the order trimethylsilyl (TMS) [1] <

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triethylsilyl (TES) [64] < t-butyldimethylsilyl (TBS) [20,000] < triisopropylsilyl (TIPS)

[700,000] < t-butyldiphenylsilyl (TBDPS) [5,000,000].57,58 Consequently, the installation of

silyl groups with varying degrees of stability onto hydroxyls with varying amounts of

substitution should allow for the selective deprotection and functionalization required. A

second major advantage to employing silyl protective groups is their ability to be cleaved

easily by a fluoride ion. Thus fluoride, typically provided in the form of tetrabutylammoniumflouride (TBAF), can be used for universal deprotection of all of the remaining silyl groups at once. In prostaglandin synthesis, however, aqueous HF is preferred due to TBAF’s tendency to give the common dehydration product, α,β-unsaturated ketone.

Because selective deprotection of the primary tail hydroxyl is essential to the success of our synthesis, the initial synthetic scheme employed TBDPS protection of the 11- and 15- secondary hydroxyl groups and TBS protection of the primary tail 20-hydroxyl. The combination of primary hydroxyl selectivity over secondary hydroxyl and the increased stability of TBDPS over TBS led us to hypothesize that this would be the safest and most effective synthetic route. However, due to the considerable steric interactions of TBDPS with our bicyclic prostamide precursor, we were unable to protect the 11-hydroxyl group.

Applying TBDPS-OTf gave only protection of the 15-hydroxyl group. Adapting the protection of the 11- and 15-hydroxyl groups to TIPS allowed for successful protection of these groups and excellent selective deprotection of the tail 20-hydroxyl group.

Thus, the overall synthetic strategy incorporates Corey’s original prostaglandin synthesis with a novel and delicate protective arrangement allowing for convergent functionalization of head and tail moieties and yielding unique prostamide analogs. The complete, successful synthesis is presented in Scheme 2.1. The overall yield of approximately 19% over sixteen steps is satisfactory for this synthesis. A detailed description and discussion of each individual step in the synthesis follows.

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Scheme 2.1 Overall synthetic scheme for the synthesis of the azido and isothiocyanato analogs with percent yields for both the R and S isomers.

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2.2.3 Synthetic Steps and Related Mechanisms

2.2.3.1 Synthesis of the ω-Chain

O O O O O TBSCl n-BuLi MeO P OMe imidazole MeO P OMe MeO P OMe THF DCM Me OH OTBS O O 10 11 Scheme 2.2 Synthesis of the phosphonate oxide to be used in the Horner-Wadsworth-Emmons reaction

The first step undertaken in the synthesis of our prostamide analogs is the creation of a phosphonate containing a hydroxyl moiety at the tail position. (Scheme 2.2) This functionality will eventually lead to a straightforward substitution at the prostamide’s C-20 position and thus give way to numerous analogs. N-Butyllithium is used to turn methyl dimethyl phosphonate into an organometallic nucleophile. Addition of ε-caprolactone establishes a new carbon-carbon bond at the lactone’s carbonyl carbon and opens the lactone to give the desired phosphonate 10. Employing ε-caprolactone as the starting material is most advantageous as it provides the desired chain length, the required C-15 position of the ketone, and the tail hydroxyl functionality all in one synthetic step. For future analogs with shorter chains β-propiolactone, γ-butyrolactone, and δ-valerolactone can be used in a comparable synthesis. Longer chains can be achieved using long-chain esters with halogen or hydroxyl functionality at the tail. The free hydroxyl tail is then protected using a silyl protecting group which is stable under the basic conditions that will generally be employed throughout the rest of the synthesis. Selective deprotection of this hydroxyl is critical to the success of this synthesis in order to functionalize only at the tail position.

Under acidic conditions, selective deprotection of a tert-butyldimethylsilyl group (TBS) should be achievable in the presence of bulkier silyl groups such as tert-butyldiphenylsilyl

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(TBDPS) and triisopropylsilyl (TIPS) groups and in the presence of TBS groups on secondary

hydroxyls.

O O

O O O O O O

MeO P OMe NaH THF O OTBS O OTBS O O 11 12

Scheme 2.3 Horner-Wadsworth-Emmons stereospecific alkylation reaction.

The prepared phosphonate 11 was treated with sodium hydride and brought together with Corey’s lactone benzoate aldehyde in a Horner-Wadsworth-Emmons reaction to give the E-enone 12 exclusively. (Scheme 2.3) Our initial synthetic attempts employed the use of a 4-phenylbenzoate protecting group at the C-11 hydroxyl as in the Corey’s

original synthesis. The 4-phenylbenzoate serves to increase the likelihood of the prostamide

intermediates achieving a solid crystalline form which could then be sent for x-ray

crystallography studies. In addition, the presence of the 4-phenylbenzoate protecting group

would only contribute aromatic proton signals to the NMR and thus would not complicate the

interpretation of the already complex NMR spectra of the prostamides. It’s enhanced UV

activity would prove an advantage during purification via TLC and flash chromatography.

Unfortunately, this group was too labile under the sodium hydroxide base used in the

Horner-Wadsworth-Emmons reaction and so a simple benzoate protecting group was

ultimately applied instead.

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2.2.3.1.1 Chiral Reduction of the 15-Keto Group

O O O O

CeCl3 NaBH O 4 O O OTBS O OTBS O OH 12 13

Separation via flash chromatography

O O O O

+ O O O OTBS O OTBS OH OH 15-S Isomer 15-R Isomer 13A 13B Scheme 2.4 Reduction of the enone to the chiral allylic alcohols.

The next step in our synthesis is the reduction of the newly established enone to the allylic alcohol 13. (Scheme 2.4) This reduction creates the final chiral center in the prostamide synthesis. Before attempting any chiral reduction methods, a simple non- stereoselective Luche reduction was carried out. The Luche reduction ensures a 1,2- chemoselective reduction as the cerium chloride coordinates to the carbonyl oxygen increasing the electrophilicity of the carbonyl carbon.59,60 The hydride then adds selectively

in a 1,2-addition rather than the competing 1,4-addition common in α,β-unsaturated

ketones. Upon separation of the resulting epimers via flash chromatography, their identity

as 15R or 15S stereocenters needed to be determined. Because these are novel compounds

with additional substitution at the tail position, optical rotation and comparison to the

literature will not be sufficient to assign the chirality. Attempts at sufficient crystallization of

the compounds were unsuccessful potentially due to the lipophilic chain tail. Mosher analysis

was decidedly the simplest and arguably the most reliable method to determine the

stereochemistry of our compounds.

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2.2.3.1.2 Mosher Analysis of the 15-Hydroxyl Stereochemistry

Mosher analysis entails derivatization of the two alcohols of unknown configuration with the two enantiomers of an auxiliary reagent, in this case Mosher’s acid:

(±)methoxytrifluoromethylphenylacetic acid (MTPA).61 The resulting diastereomeric esters are then subjected to proton NMR and differences in chemical shifts are measured. The differences in a proton’s chemical shift upfield or downfield indicates the orientation of those protons around the enantiomeric center coupled to the chiral auxiliary.62

H O H O

CF CF L2 R 3 L2 S 3 O O L1 L1 Ph OMe MeO Ph

Shielding

OCF3 OCF3

e MeO Shielding Ph OM Ph L1 L2 L1 L2 R-Mosher S-Mosher

Figure 2.14 The preferred orientation of R and S Mosher esters coupled to a generic chiral center shown as standard and Newman projections. The arrows indicate the shielding effect on the rest of the compound by the Mosher ester phenyl group.

According to Mosher, the MTPA-esters in solution exist mainly in a single conformation where the bond between the central mosher carbon and the trifluromethyl group is nearly syn-periplanar to the carbonyl bond of the ester.61 (Figure 2.14) The ester

adopts the standard s-trans arrangement allowing for both the methine proton of the

secondary alcohol and the trifluoromethyl to be syn-coperiplanar with the carbonyl. In this orientation the protons resident on one side of the secondary alcohol experience shielding

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effects from the phenyl substituent. For example, introduction of the R-Mosher ester in

Figure 2.14 gives shielding of the protons on the L1 side of the alcohol and NMR signals

correlating to these protons would be expected to shift upfield in comparison with the S-

Mosher. Although this is not the only conformation achievable by these freely rotating

diastereomers, the model correlates to the average shielding effects observed under NMR.

Thus, lower temperature NMR yields greater differences in shifting of the spectral signals.

Fortunately, we have established two isomers that are separable by flash chromatography, and thus, we have the ability to confirm our Mosher analysis results for one isomer with those of the other. The isomers have been arbitrarily assigned as “Isomer

A” – the less polar isomer with lower retention time on silica, and “Isomer B” – the more polar isomer with higher retention time on silica. With this in mind, each isomer was isolated and coupled separately to each of the Mosher chlorides. (Figure 2.15)

O O (R or S)

O O OTBS OH 13 O O O “B”,”S” O “B”,”R” (R) or (S)-Mosher Acid Chloride DMAP O TEA O OTBS O O OTBS DCM O O O O CF 3 CF3 Ph OMe MeO Ph “Isomer A” O “A”,”S” “A”,”R” O O “Isomer B” O

O O OTBS O O OTBS A – Less Polar (Higher Rf) B – More Polar (Lower Rf) O O O O CF3 CF3 Ph OMe MeO Ph

Figure 2.15 Coupling and assignment of the Mosher acid chloride to the alcohols of unknown configuration.

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Extreme care must be taken in this analysis as conversion of the Mosher acid chloride to the

Mosher ester reverses the stereochemical assignment of the auxiliary’s chiral center. While

the configuration remains entirely unchanged, the substitution of an oxygen atom for a

chlorine atom rearranges the order in which the substituents are assigned according to the

Cahn-Ingold-Prelog priority rules for assigning stereochemistry to chiral centers. (Figure

2.16) To avoid confusion, within the confines of this study we will employ the Mosher

analysis in such a way that the “R/S-mosher” refers to the R/S mosher acid chloride before

it is coupled to the alcohol and not to the R/S ester product. For example, the R-mosher of

isomer “A” denotes the alcohol with the higher retention factor (Rf) value coupled to the R-

mosher acid chloride. This becomes most important in the final assignment of the chiral

alcohols.

Acid Chloride Ester

O O 2 3 3 2 Cl OR* F3C F3C Ph OMe Ph OMe 4 1 4 1 "R" "S"

O O 3 2 2 3 Cl OR* F3C F3C MeO Ph MeO Ph 1 4 1 4 "S" "R" Figure 2.16 Inversion of the stereochemical assignment under the Cahn-Ingold-Prelog priority rules when the Mosher acid chloride reacts with the chiral alcohol to give the diastereomeric ester.

The 1H NMR spectra of the resulting diastereomers were obtained, and following assignment of the protons, the proton shifts of isomers “A” and “B” under the influence of the R and S mosher esters were compared. The results are represented spectrally in Figure

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2.17 and are quantitatively presented in Figure 2.18. Three specific protons were chosen to illustrate the effective shifts in spectra due to their ease of identification and proximity to the chiral center. While nearly all of the protons will experience a shift in their spectral signal in the direction dictated by the orientation of the mosher ester, the greatest effect will be observed on the signals given by protons nearest the diastereomeric center.

Conveniently, the closest protons on the cyclopentyl side of the stereocenter are the easily identifiable alkene protons, each as a doublet of doublets coupled to each other at around

5.6 ppm. These protons exhibit the greatest change in chemical shift. Unfortunately, the protons directly on the other side of the chiral center fall in the range of standard alkyl protons and are difficult to distinguish via NMR. The protons nearest the protected hydroxyl tail are the most easily identifiable as a triplet at around 3.5 ppm. Because these protons

Isomer A coupled to R-Mosher acid

Isomer A coupled to S-Mosher acid O O

O 1 3 O OTBS 2 Isomer B coupled to R-Mosher acid OMTPA

Isomer B coupled to S-Mosher acid

Figure 2.17 Set of NMRs for the four Mosher esters synthesized with analyzed protons assigned. (1) Corresponds to the alkene proton adjacent to the hydroxyl and the red circled proton signal, (2) corresponds to the allylic alkene proton and the green circled proton signal, and (3) corresponds to the tail methylene protons adjacent to the protect hydroxyl and the blue circled proton signal.

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are more distal from the stereocenter, the effective shift will be small. However, it should be in the opposite direction to the alkene protons. Ultimately, it is the direction of the shift and thus the sign on the Δ δSR and not the magnitude of the shift that is important. Consistency

in the direction of proton shifts of all protons on either side of the stereocenter is essential.

Additionally, the effective shifting of these distal proton signals may indicate a propensity of

the terminal TBS-protected hydroxyl to loop back around and approach the 15-hydroxyl.

Figure 2.17 shows the overall effect of the mosher esters on the highlighted proton

signals of the two isomers. The alkene protons (1,2) of isomer “A” are shifted significantly

downfield when coupled to the R-mosher acid chloride compared with the upfield shift when

coupled to the S-mosher acid chloride. Within the same isomer, the distal tail protons (3)

are shifted by a lesser degree in the opposite directions. R-mosher acid chloride incites an

upfield shift, while coupling to the S-mosher acid chloride gives a downfield shift. The proton

shifts observed with mosher acids coupled to the “B” isomer are entirely opposite to these

“A” isomer results. The R-mosher acid chloride gives an upfield shift to the alkene protons

(1,2) and a downfield shift to the distal tail protons (3), while the S-mosher acid chloride

gives a downfield shift to the alkene protons (1,2) and an upfield shift to the distal tail

protons (3). Figure 2.26 quantifies the resulting shift values and gives the Δ δSR for each of

the highlighted protons. Based on the direction of these shifts and the knowing the

stereochemistry of the mosher esters they are coupled to, we can determine the absolute

stereochemistry of the 15-hydroxyls in the two isomers “A” and “B”.61-64

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A δS –AS (ppm) δR –AR (ppm) Δ δSR (ppm) Δ δSR (Hz) 0 5.440 5.426 0.018 9 O O 1 5.730 5.591 0.139 69.5 2 5.664 5.560 0.104 52.0 3 O 1 3 3.555 3.577 -0.022 -11.0 O OTBS 2 0 OMTPA B δS –BS (ppm) δR –BR (ppm) Δ δSR (ppm) Δ δSR (Hz) 0 5.415 5.422 -0.007 -3.5 1 5.631 5.735 -0.104 -52.0 2 5.513 5.628 -0.115 -57.5 3 3.544 3.524 0.020 10.0

H H (2) H (3) SR = Δδ >0 CH CH(CLB) (CH2)5OTBS MTPAO MTPAO MTPAO = (1) (CH2)5OTBS (1) CH CH(CLB) ΔδSR<0 (3) (2) “A” = S “B” = R Important: R and S in ΔδSR denote stereochemistry of the acid chloride before coupling to the alcohol

Figure 2.18 Analysis of the Mosher esters with corresponding proton shifts tabulated and a pictorial representation of the Mosher conventional assignment under the Cahn-Ingold-Prelog priority rules.

According to our NMR results, for isomer “A” the Δ δSR for the alkene protons are positive and the Δ δSR for the tail protons are negative. Using the conventions of mosher

analysis shown pictorially in Figure 2.18, the alkene protons are assigned to the open blue

circle and the tail protons are assigned to the solid blue circle.62 Following the Cahn-Ingold-

Prelog priority rules for assigning stereochemistry, isomer “A” is the S isomer. Following

mosher analysis conventions a second time leads us to the conclusion that isomer “B” is the

R isomer. This is in agreement with previous assignments done on prostaglandin

intermediates with a straight chain tail where the less polar compound was determined to

be the 15-S secondary alcohol and the more polar compound was found to be 15-R.65

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2.2.3.1.3 Attempts at Chiral Reduction of the 15-Keto Group

H Ph O Ph RS OH R R Ph + S L O N RL O B - + B H N Ph RS RL B- R-oxazabolidine H H

Figure 2.19 Mechanism of chiral induction during Corey’s stereoselective oxazaborolidine reduction

Now that we have determined which isomer has the R configuration and which has the S configuration following the enone reduction, we can quantify the results of our attempts at chiral reduction. The non-stereoselective Luche reduction yielded an enantiomeric excess of approximately 30% in favor of the S isomer. To see if enantioselectivity could be achieved using the well-established methods employed by

Corey’s group, Corey’s oxazaborolidine catalyst (CBS) was used along with borane in either

THF or DMS.66 Using one of Corey’s optically pure oxazaborolidine catalysts in the standard prostaglandin synthesis gives approximately 80% enantiomeric excess of the corresponding chiral alcohol.67 According to the literature, R-oxazaborolidine provides mainly the 15S native conformation, and S-oxazaborolidine gives the 15R conformation selectively. The proposed sterically driven mechanism for this stereoselectivity involves first the borane binding to the nitrogen of the R-oxazaborolidine and the coordination of the R- oxazaborolidine boron to the carbonyl oxygen of the enone. (Figure 2.19) The intermediate adopts a chair-like configuration in which the smaller alkyl group of the enone aligns with the methyl of the R-oxazaborolidine and the larger group faces away from the steric bulk of the R-oxazaborolidine methyl and pyrrolidine ring. The borane is able to then deliver the hydride to the forward facing side of the carbonyl giving selectively the S-chiral alcohol. The

S-oxazaborolidine contains a pyrrolidine ring facing the opposite direction and arranges its

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methyl along side it during the reaction. Thus the enone aligns in the reverse direction and

the hydride will be delivered to the opposing face giving the R-chiral alcohol selectively.

Unfortunately, after several attempts employing the R-oxazaborolidine catalyst for

the reduction of our functionalized analogs, we achieved only 30-39% enantiomeric excess

of the desired S-chiral alcohol. In fact, we achieved levels of enantiomeric excess very

similar to those observed during the Luche reduction. Our enantiomeric excess did not

approach that of Corey’s synthesis, perhaps due to the relatively large TBS protecting group resident on the tail hydroxyl. This group may give enough steric bulk to rival that of the bicyclic group. Without a great disparity between the size of the structure to “the left” and

to “the right” of the ketone, the selectivity of the reaction will not be optimal. Indeed because the enantiomeric excess is almost identical to that of the Luche reduction, there is

also a possibility that the oxazaborolidine is not capable of coordinating in any meaningful

way to our functionalized analogs. The greater steric bulk of our analogs compared to that of the straight chain tail of the prostaglandins may account for this disparity. In agreement

with the Mosher analysis, these results may also suggest that tail hydrocarbon chain curves

around back towards the 15-hydroxyl portion of the molecule. This orientation could further

block the coordination of the oxazaborolidine to the enone during the attempted chiral

reduction.

Before moving on, a different chiral reduction was attempted using S-(1,1’)- binaphthalene-2,2’-diol (S-BINOL). Like the oxazaborolidine, this chiral auxiliary has found

use in the chiral reduction of prostaglandin 15-keto intermediates to the 15S-alcohol.68,69

BINOL forms a complex with lithium aluminum hydride which then delivers a hydride to the

carbonyl carbon giving the 1,2-chemoselective reduction. (Figure 2.20) The stereochemical

orientation of the BINOL determines the stereoselective outcome. In standard prostaglandin

synthesis, S-BINOL gives the desired 15S-alcohol and R-BINOL gives the unnatural 15-

alcohol. Unfortunately, our analogs again defied convention and would not give substantially

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greater enantiomeric excess beyond that of the Luche control reaction. Once more, steric

bulk of the TBS protecting group could be the complicating factor that keeps the chiral

reducing agent from being able to prefer one orientation over the other.

Figure 2.20 Mechanism of chiral induction during stereoselective BINOL reduction

Table 2.1 Stereoselectivity of the chiral and achiral reductions of the prostamide enone Reaction Attempted % S Isomer % R Isomer % Enantiomeric Excess

R-CBS, BH3-THF (1) 65 34 30

R-CBS, BH3-THF (2) 69 30 39

R-CBS, BH3-DMS 65 34 30

S-Binol 65 34 31

Luche (1) 67 32 34

Luche (2) 64 35 29

Table 2.1 gives a summary of the stereochemical outcomes of the stereoselective and non-stereoselective reductions attempted in our lab. In the interest of time and seizing the opportunity to double our library of compounds, we decided to forgo the

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enantioselective reduction and isolate both the R and S isomers for further synthesis.

Because the isomers were fairly easily separated at this stage, and their absolute

configuration successfully determined by the Mosher analysis, the remainder of the

synthesis was done in parallel. This strategy negates the possibility that the isomer

retention factor values could merge or even reverse throughout the synthesis, making

purification difficult or impossible and requiring a second analysis of the 15-hydroxyl

stereochemistry.

Hydrolysis of the benzoyl protecting group using aqueous base was the next step in

the total synthesis of our prostamide compounds. (Scheme 2.5) While overall a very basic

synthetic step, the isolation of the product proved to be more challenging than anticipated.

Poor solubility of our compounds in MeOH, and opening of the lactone ring to form a

potassium salt complicated our initial attempts and left us with poor yields. However,

dissolving the starting material in THF before addition to the reaction mixture

(THF:MeOH,1:10), ensuring 1.0 equivalent of potassium carbonate, and, most importantly,

carefully working-up with saturated ammonium chloride gave excellent yields of the lactone

diol 14. Additionally, as previously proposed, we observed a reversal of the R and S

isomers’ Rf values upon removal of the benzoyl group. This exchange gave added support to the approach of proceeding with the R and S isomers in parallel throughout the rest of the

synthesis.

Scheme 2.5 Hydrolysis of the benzoyl protection and silyl protection of the diol.

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To ensure a substantial difference between the strength of the protecting groups on

the two secondary hydroxyls and the tail hydroxyl, the bulky tert-butyldiphenylsilyl was

selected as protecting groups for the secondary hydroxyl. In general, bulkier silyl groups are

more stable to acid hydrolysis. Additionally, primary silyl ethers are more labile than their

secondary equivalents. In practice, the tert-butyldiphenylsilyl turned out to be too bulky for

the protection of the 11-hydroxyl. After days of reaction with TBDPS triflate, more reactive

than TBDPS chloride, at elevated temperature to 50ºC, TBDPS protection only occurred at

the 15-hydroxyl position (81% yield). To compensate, the medium-sized triisopropylsilyl

(TIPS) protective group was employed and successfully protected both secondary hydroxyls

(92% yield). The additive effects of bulkier TIPS over the TBS group and the more stable

secondary positions of the 11- and 15-hydroxyl groups should allow for the facile selective

deprotection of the tail hydroxyl. In addition, due to the difficulty encountered on

introduction of the TBDPS group at the 11-hydroxyl, the deprotection of this group within

the final steps of the synthesis would most likely be quite challenging. Adopting a silyl

protecting group that is more easily introduced gave an indication that it will likely be more

easily removed as well.55

2.2.3.2 Synthesis of the α-Chain

The next stage in the synthesis was the introduction of the head chain appendage.

The lactone 15 was reduced to the lactol 16 using diisobutylaluminum hydride (Dibal-H) in

preparation for a ring opening Wittig reaction. (Scheme 2.6) Without this partial reductive

step, the Wittig reagent would not be able to force the ring opening. Following the

conventions of Wittig reactions discussed earlier, the unstable ylide, (4-Carboxybutyl) triphenylphosphonium bromide, added across the electrophilic lactol carbon to open the ring and give the C5-C6 Z-alkene 17 exclusively.

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Scheme 2.6 Reduction of the lactone to the lactol and Wittig reaction.

With both the long chain appendages installed, focus was shifted to the funtionalization of the head and tail moieties. This key intermediate 17 was the common precursor to which modifications of the α and ω substituents could be made. The opportunity now arose to either establish first a common head modification and vary the tail or introduce a common tail group and modify the head attachments, the goal being to accomplish a small set of analogs with diverse head substituents and a small set with differing tail substituents. Following feed-back from biological testing, the set can be combined to achieve the optimal combination of head and tail group moieties. In our case, we chose to institute a common, covalent-binding tail group and make several head group modifications. With this in mind, reaction of the tail hydroxyl group to provide azido and isothiocyanato functionalities was next to be undertaken. Before this conversion could be achieved, the other free hydroxyls must be masked and the tail hydroxyl must be selectively deprotected.

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2.2.3.3 Functionalization of the ω-Tail

Scheme 2.7 Methyl esterification, oxidation of the 9-hydroxyl group, and selective deprotection of the 20-hydroxyl group

The free carboxylic acid 17 was protected by conversion to the methyl ester 18 using trimethylsilyldiazomethane (TMS-CH2N2) in methanol. (Scheme 2.7) The mechanism inferred for this reaction begins with the deprotonation of the carboxylic acid by the diazomethane.70 Methanol then donates a proton back to the carboxylic acid before reacting with the silyl group. TMS-protected methanol is produced along with the carboxylic acid and diazomethane. From here, the methylene deprotonates the acid a second time and the carboxylate then reacts with the methyl to produce the methyl ester and release nitrogen gas in the process. (Figure 2.21)

Figure 2.21 Mechanism of methyl ester formation from carboxylic acid by treatment trimethylsilyldiazomethane in methanol.

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The 9S-hydroxyl on the ring was then oxidized to the ketone 19 using a Dess-Martin

oxidation. Dess-Martin Periodinane (DMP) is a hypervalent iodine compound that allows for

a very mild and chemoselective oxidation of alcohols to ketones.71,72 The terminal TBS

protected hydroxyl then needed be selectively deprotected in the presence of the secondary

TIPS protected hydroxyl groups. The best method for this conversion was determined to be

a mixture of acetic acid, tetrahydrofuran (THF), and water (AcOH:THF:H2O – 3:1:1). Under

these mild conditions, deprotection of the silyl groups occurs slowly and selectively to cleave

only the most labile terminal TBS group. 1% HCl is also very effective in deprotecting the

primary TBS group, however because of its strength some di-deprotection occurs after only

ten minutes while substantial starting material is still present.

Figure 2.22 Key intermediate from which further tail modifications can easily be made.

At this critical point we had established the means to functionalize the tail with any

number of substitutions. Conversion to a mesylate, tosylate, or triflate would allow for SN2

reaction with an array of nucleophiles. (Figure 2.22) Our initial synthesis focused only on

the azido and isothiocyanato groups to serve as covalent probes in receptor sites. A

Mitsunobu-type reaction using zinc azide/bis-pyridine, diisopropyl azodicarboxylate (DIAD),

and triphenylphosphine (PPh3) provided the subsequent conversion of the tail hydroxyl 20 to the azide 21. (Scheme 2.8) As with a standard Mitsunobu reaction, the first step in the

reaction mechanism is the nucleophilic attack by the PPh3 upon one of the azo nitrogens of

the DIAD. The electron density of this betaine, now shifted to the other nitrogen, allows for

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it to react with the Zn azide complex and release an azide ion. In the standard Mitsunobu

reaction, the nitrogen would instead be protonated by the acidic reagent that becomes the

reaction’s nucleophile. The alcohol contributes a proton to the first nitrogen, and in return

forms a bond with the PPh3. Nucleophilic attack by the azide ion releases triphenylphosphine

oxide and forms the necessary azido compound.

Scheme 2.8 Modified Mitsunobu and reduction of the ring ketone.

2.2.3.4 Functionalization of the α-Head Amide

With the tail modification in place, focus was shifted to the head functionality. Amide coupling was attempted several times using different methods and under a variety of conditions. Reactions attempted included enzymatic amidation using a variety of lipases, standard peptide-like coupling using 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide

(EDCI) and 1,1'-carbonyldiimidazole (CDI), amidation by formation of an acid chloride using

oxalyl chloride and of an acyl cyanide using sodium cyanide, and finally a mixed anhydride

amidation using isobutyl chloroformate. However, each reaction attempted led to a large

portion of the α,β-unsaturated ketone side product by elimination of the 11-hydroxyl. It was

clear that the 9-keto moiety would have to be protected during this step of the synthesis.

Protection by establishing dimethoxy (ketal) functionality at the C-9 position was attempted

numerous times. Such a protection would allow for selective protection of the ketone and

facile deprotection back to the ketone under acidic condition similar to those used to

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deprotect the silyl groups at the end of the synthesis. Unfortunately, this reaction turned

out to be rather difficult, giving a good yield only once out of eight attempts, and causing a

substantial amount of TIPS deprotection. Because this protection turned out to be unnecessarily difficult, we decided instead to reduce the ketone back to the alcohol simply to lessen the propensity of the compound to eliminate to the conjugated form. In this way,

our synthetic strategy was adapted such that the conversion of the 9-hydroxyl 18 to the ketone 19 was purely intended as a type of “protection” against conversion of the alcohol to the azide during the Mitsunobu step. This was preferable to the establishment of yet another silyl protecting group that would require selective deprotection prior to oxidation to the ketone. Thus the 9-keto group in 21 is reduced back to the diasteriomeric 9-hydroxyl

R/S-22 using the same Luche conditions as in the previous reduction.

Scheme 2.9 Hydrolysis of the methyl ester, amidation, and oxidation of the 9-OH hydroxyl group

The methyl ester 22 was hydrolyzed to the acid 23 using a hydrolysis in aqueous

base. (Scheme 2.9) Before efforts to synthesize the amide were undertaken, the ethanolamine reagents had to be protected. This was required, not because the alcohol may react to give the ester, as the amine is more reactive, but because the oxidation 24 of the

9-hydroxyl back to ketone 25 is not chemoselective. Our effort to use Dess-Martin oxidation produced only the terminal aldehyde. A separate attempt was made to protect this

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ethanolamide alcohol with a silyl ether group prior to oxidation, but surprisingly the 9- hydroxyl was selectively protected over the terminal head alcohol. Thus, the ethanolamine analogs were protected with TBS groups prior to amide coupling. In order to prevent TBS protection at the amine, the amine was first selectively protected with benzyl chloroformate, then the alcohol was protected with TBSCl, and finally the amine was deprotected by a hydrogenation with hydrogen and palladium. (Scheme 2.10)

Scheme 2.10 Synthesis of the TBS protected ethanolamine

Figure 2.23 Mechanism of amide coupling by 1,1'-Carbonyldiimidazole (CDI)

As discussed previously, several coupling reagents were used in an attempt to

provide the amide from both the methyl ester and the free carboxylic acid. Of these

reactions, the amidation coupling using CDI was determined to be the easiest and highest yielding. CDI initiates the reaction by deprotonating the acid and serving as an electrophile for attack by the carboxylate ion and releasing imidazole. (Figure 2.23) The imidazole then

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reacts with the acyl carboxy imidazole to give the activated acylimidazole. After allowing for

this conversion to take place, addition of the amine will give a nucleophilic attack at the

carbonyl and release a second molecule of imidazole. This robust reaction was effective

towards all of the varied amines added, providing the desired amide 24. With this critical step complete, the 9-hydroxyl moiety was oxidized under the standard Dess-Martin conditions back to the ketone 25.

The final step in the synthesis of the initial set of azido analogs AM7637, AM7638,

AM7651, AM7652 was universal deprotection of the silyl groups to give 26. (Scheme 2.11)

This was accomplished using 48% hydrogen fluoride over 24 hours in a Teflon tube. The isothiocyanato compounds needed one more step in which the azido group was transformed to the isothiocyanate when treated with carbon disulfide. The approximate overall yield for the synthesis of these compounds from Corey’s lactone benzoate aldehyde starting material was approximately 19%. Given the substantial number of steps in synthesis (16 steps), this yield is quite acceptable. The synthesis has proven robust and the results have been repeated several times within our lab.

Scheme 2.11 Final universal deprotection and conversion of the azido to the isothiocyanato group.

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2.2.3.5 Synthesis of the Non-Functionalized ω-Chain and Glycerol Ester Analogs

Subsequent syntheses were carried out both to optimize conditions and to complete

the remaining proposed analogs. The final deprotection step of the initial synthesis proved

to be challenging and provided relatively low yields due to the stability of the secondary

TIPS at the 11-hydroxyl position. Under these hydrogen fluoride conditions, this hydroxyl

group was the most difficult to deprotect. The full synthesis was repeated using TBS at this

position and 1%HCl in 95% MeOH for the final universal deprotection, leading to significantly higher yields. Thus, TBS is a viable alternative for protection of the 11- hydroxyl. This group would theoretically give slightly less selectivity during deprotection of the TBS protected 20-hydroxyl, however, in practice, the selectivity was excellent and the final deprotection benefited greatly from the greater reactivity of the TBS group compared to the TIPS.

Scheme 2.12 Synthesis of the analogs with straight carbon chain tail

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The analogs with the straight terminal alkyl chain were less complex to synthesize, requiring only six steps in a method similar to standard prostaglandin synthesis. (Scheme

2.12) Protection of the diol 28 with TBS groups followed by nonstereospecific reduction of the lactone to the lactol prepares the compound for a standard Wittig, giving Z-alkene 31.

Amide coupling, Dess-Martin oxidation, and cleavage of the TBS protecting groups gives the

final AM7648, AM7649, and AM7650 compounds.

Synthesis of the 2-glycerol analog began with the free acid 35 coupling to the

butyrate protected glycerol. (Scheme 2.13) Oxidation of the free hydroxy on the ring was

followed by the deprotection of both silyl protecting groups. Finally, the 2-glycerol was

unmasked by an esterase cleavage of the protective butyrates to reveal the desired 2-

glycerol analog 39.

Scheme 2.13 Synthesis of the prostaglandin glycerol ester analog

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36. Serhan, C. N.; Yacoubian, S.; Yang, R., Anti-inflammatory and proresolving lipid mediators. Annu. Rev. Pathol.: Mech. Dis. 2008, 3, 279.

37. Hamberg, M., Isolation and Structure of Two Prostaglandin Endoperoxides that Cause Platelet Aggregation. Proc. Natl. Acad. Sci. U. S. A. 1974, 71, 345.

38. Peng, S.; Okeley, N. M.; Tsai, A.-L.; Wu, G.; Kulmacz, R. J.; van der Donk, W. A., Structural Characterization of a Pentadienyl Radical Intermediate Formed during Catalysis by Prostaglandin H Synthase-2. J. Am. Chem. Soc. 2001, 123, 3609.

39. Serhan, C., Lipoxins and aspirin-triggered 15-epi-lipoxins are the first lipid mediators of endogenous anti-inflammation and resolution. Prostaglandins, Leukotrienes Essent. Fatty Acids 2005, 73, 141.

40. Romano, M., Lipoxin and aspirin-triggered lipoxins. TheScientificWorldJournal 2010, 10, 1048.

41. Serhan, C. N. M., J. F.; Petasis, N. A.; Akritopoulau-Zanze, I.; Papayianni, A.; Brady, H. R.; Colgan, S. P.; Madara, J. L. Biochemistry 1995, 1995, 14609.

42. Perera, S. K.; Fedor, L. R., Acid- and base-catalyzed dehydration of prostaglandin E2 to prostaglandin A2 and general-base-catalyzed isomerization of prostaglandin A2 to prostaglandin B2. J. Am. Chem. Soc. 1979, 101, 7390.

43. Bindra, J. S.; Bindra, R. Prostaglandin Synthesis; Academic Press: New York, 1977.

44. Mitra, A. The Synthesis of Prostaglandins; Wiley-Interscience: New York, 1977.

45. New Synthetic Routes to Prostaglandins and Thromboxanes; Academic Press: San Diego, 1982.

46. Caton, M. P. L., A survey of novel and useful reactions discovered through research on prostaglandins. Tetrahedron 1979, 35, 2705.

47. Nicolaou, K. C.; Gasic, G. P.; Barnette, W. E., Synthesis and biological properties of prostaglandin endoperoxides, thromboxanes and prostacyclins. Angew Chem Int Ed Engl 1978, 17, 293.

48. Newton, R. F.; Roberts, S. M., Steric control in prostaglandin synthesis involving bicyclic and tricyclic intermediates. Tetrahedron 1980, 36, 2163.

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49. Noyori, R.; Suzuki, M., New synthetic methods. (49). Prostaglandin syntheses by three- component coupling. Angew. Chem. 1984, 96, 854.

50. Corey, E. J.; Weinshenker, N. M.; Schaaf, T. K.; Huber, W., Stereo-controlled synthesis of dl-prostaglandins F2α and E2. J. Amer. Chem. Soc. 1969, 91, 5675.

51. Corey, E. J.; Schaaf, T. K.; Huber, W.; Koelliker, U.; Weinshenker, N. M., Total synthesis of prostaglandins F2α and E2 as the naturally occurring forms. J. Amer. Chem. Soc. 1970, 92, 397.

52. Corey, E. J.; Noyori, R.; Schaaf, T. K., Total synthesis of prostaglandins F1α, E1, F2α, and E2 (natural forms) from a common synthetic intermediate. J. Amer. Chem. Soc. 1970, 92, 2586.

53. Corey, E. J., Total synthesis of prostaglandins. Ann. N. Y. Acad. Sci. 1971, 180, 24.

54. Corey, E. J. The Logic of Chemical Synthesis; John Wiley & Sons: New York, 1989.

55. Greene, T. W.; Wuts, P. G. M. In Protective Groups in Organic Synthesis; 3 ed.; Greene, T. W., Wuts, P. G. M., Eds.; John Wiley & Sons: New York, 1999, p 113.

56. van Look, G.; Simchen, G.; Heberle, J. Silylating Agents; 2 ed.; Fluka Chemie AG: Buchs, Switzerland, 1995.

57. Shimizu, N.; Takesue, N.; Yasuhara, S.; Inazu, T., Prediction of structural effects of trialkylsilyl groups on reactivity toward nucleophilic displacement at silicon. Chem. Lett. 1993, 1807.

58. Shimizu, N.; Takesue, N.; Yamamoto, A.; Tsutsumi, T.; Yasuhara, S.; Tsuno, Y., A quantitative scale for the structural effect on reactivity toward nucleophilic displacement at silicon. Chem. Lett. 1992, 1263.

59. Luche, J. L., Lanthanides in organic chemistry. 1. Selective 1,2 reductions of conjugated ketones. J. Am. Chem. Soc. 1978, 100, 2226.

60. Gemal, A. L.; Luche, J. L., Lanthanoids in organic synthesis. 6. Reduction of α-enones by sodium borohydride in the presence of lanthanoid chlorides: synthetic and mechanistic aspects. J. Am. Chem. Soc. 1981, 103, 5454.

61. Dale, J. A.; Mosher, H. S., Nuclear magnetic resonance enantiomer regents. Configurational correlations via nuclear magnetic resonance chemical shifts of diastereomeric mandelate, O-methylmandelate, and .alpha.-methoxy-.alpha.- trifluoromethylphenylacetate (MTPA) esters. J. Am. Chem. Soc. 1973, 95, 512.

62. Hoye, T. R.; Jeffrey, C. S.; Shao, F., Mosher ester analysis for the determination of absolute configuration of stereogenic (chiral) carbinol carbons. Nat. Protoc. 2007, 2, 2451.

63. Latypov, S., Determination of the absolute stereochemistry of alcohols and amines by NMR of the group directly linked to the chiral derivatizing reagent. Tetrahedron 2001, 57, 2231.

64. Seco, J., A practical guide for the assignment of the absolute configuration of alcohols, amines and carboxylic acids by NMR. Tetrahedron: Asymmetry 2001, 12, 2915.

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65. Fleming, I.; Winter, S. B. D., Stereocontrol in organic synthesis using silicon-containing compounds. A formal synthesis of prostaglandins controlling the stereochemistry at C-15 using a silyl-to-hydroxy conversion following a stereochemically convergent synthesis of an allylsilane. J. Chem. Soc., Perkin Trans. 1 1998, 2687.

66. Galatsis, P.; John Wiley & Sons, Inc.: 2007, p 2.

67. Corey, E. J.; Helal, C. J., Reduction of carbonyl compounds with chiral oxazaborolidine catalysts: A new paradigm for enantioselective catalysis and a powerful new synthetic method. Angew. Chem., Int. Ed. 1998, 37, 1986.

68. Noyori, R.; Tomino, I.; Tanimoto, Y.; Nishizawa, M., Asymmetric synthesis via axially dissymmetric molecules. 6. Rational designing of efficient chiral reducing agents. Highly enantioselective reduction of aromatic ketones by binaphthol-modified lithium aluminum hydride reagents. J. Am. Chem. Soc. 1984, 106, 6709.

69. Noyori, R.; Tomino, I.; Yamada, M.; Nishizawa, M., Asymmetric synthesis via axially dissymmetric molecules. 7. Synthetic applications of the enantioselective reduction by binaphthol-modified lithium aluminum hydride reagents. J. Am. Chem. Soc. 1984, 106, 6717.

70. Kuhnel, E.; Laffan, D. D. P.; Lloyd-Jones, G. C.; Martinez, D. C. T.; Shepperson, I. R.; Slaughter, J. L., Mechanism of methyl esterification of carboxylic acids by trimethylsilyldiazomethane. Angew Chem Int Ed Engl 2007, 46, 7075.

71. Dess, D. B.; Martin, J. C., Readily accessible 12-I-5 oxidant for the conversion of primary and secondary alcohols to aldehydes and ketones. J. Org. Chem. 1983, 48, 4155.

72. Dess, D. B.; Martin, J. C., A useful 12-I-5 triacetoxyperiodinane (the Dess-Martin periodinane) for the selective oxidation of primary or secondary alcohols and a variety of related 12-I-5 species. J. Am. Chem. Soc. 1991, 113, 7277.

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Chapter 3: Evaluation of Prostamides by Receptor Screening and as Lipid Mediators in Inflammation and Immunomodulation

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3.1 Biological Screening of Prostamides on Established and Orphan Receptors

The initial aim of this study was to provide compounds for screening against a wide

range of receptors for the purpose of identifying a novel interaction. Receptor screening

serves as a fundamental preliminary investigation when exploring the biological activity of

novel compounds. The first four compounds synthesized (AM7637, AM7638, AM7645,

AM7646) were sent, along with a commercial PGE2-EA standard, for the initial receptor screening at the Roth Lab at the University of North Carolina – Chapel Hill as part of the

National Institute of Mental Health (NIMH) Psychoactive Drug Screening Program.† (Figure

3.1) Within this program, compounds are subjected to a cache of primary assays designed

to identify a subset of potential receptors or transporters for which the test compounds

display affinity. Once identified, these compounds undergo a secondary round of screening

on these same receptors or transporters in which concentration dependent curves are

established and Ki values are determined.

Figure 3.1 Compounds prepared and sent for receptor screening

The results of these screenings indicated that our prostamide compounds were inactive at adrenergic, cannabinoid, dopamine, gaba, histamine, muscarinic, serotonin, sigma, opioid and cholecystokinin receptors.† (Table 3.1) Inactivity in the primary

† The receptor screening was done by Gabriela Salazar, Ryan Whaley, Thomas Mangano, Alice Jiang, Sandy Hufeisen, and Xi-Ping Wuang in Dr Roth’s Lab at UNC-Chapel Hill

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screening is defined by compounds having less than 50% inhibition of radioligand binding at

10 µM. Additionally, these AM compounds did not show affinity for the transporter proteins,

norepinephrine transporter and serotonin transporter receptors. However, the azido

prostamides, AM7637 and AM7638, did show some affinity for the dopamine transporter

receptor (DAT). AM7638, the 15-R hydroxy 20-azido analog, showed the greatest affinity for

DAT within the primary screen giving 86% displacement of [3H]WIN35428 from the transporter protein at 10 µM.

Table 3.1 Primary Radioligand Binding Assay Results at 10 µM for 46 of the well-known receptors. Highlighted cells indicate a significant radioligand inhibition exceeding 50%. Given as mean % inhibition (n=4)

Receptor PGE2-EA AM7637 AM7638 AM7645 AM7646 Tested Final Azide S Final Azide R Final NCS R Final NCS S Serotonin 5ht1a 8.1 11.8 12.2 9.3 15.2 5ht1b 36.5 42.1 33.4 24.5 29.6 5ht1d 10.3 -0.7 22.0 -3.0 2.6 5ht1e 2.8 5.7 2.6 -1.1 1.7 5ht2a 12.6 14.8 19.1 11.7 23.4 5ht2b 11.3 -13.3 -2.1 -2.3 18.0 5ht2c 42.6 26.0 32.2 26.9 1.5 5ht3 17.5 -0.8 0.5 1.5 0.7 5ht5a 19.9 41.0 42.2 27.0 28.9 5ht6 6.2 -0.6 2.3 1.6 10.8 5ht7 5.4 7.3 -0.2 -6.2 2.7 Adrenergic Alpha1A 11.1 -5.9 -4.1 -0.6 -12.2 Alpha1B -12.8 -14.1 -4.8 -18.1 -7.4 Alpha1D 6.1 -3.5 -0.6 15.9 -3.3 Alpha2A -4.9 -2.2 -3.9 42.5 37.1 Alpha2B -0.5 8.1 0.4 -2.9 -0.1 Alpha2C -6.0 17.4 19.8 18.2 21.2 Beta1 -1.1 -1.0 -0.6 -0.2 3.9 Beta2 -8.4 3.0 -3.1 7.3 -1.4 Beta3 -1.4 21.0 16.5 15.1 -0.3 Benzodiazepine BZP Rat Brain 35.6 29.3 31.4 34.0 33.3 Site Cannabinoid CB1 -14.7 -17.5 -8.4 -6.8 -4.3 CB2 (secondary) (secondary) (secondary) (secondary) (secondary)

(contd on next page)

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Dopamine D1 26.8 32.8 29.5 32.2 3.9 D2 -8.7 -3.5 -1.6 -5.5 4.0 D3 -1.1 -5.3 2.3 14.1 17.1 D4 1.7 16.2 -6.6 -6.9 -12.5 D5 16.7 20.6 64.2 47.9 45.2 Transporter DAT 59.3 77.6 86.3 19.9 22.4 NET 12.6 11.5 3.7 2.7 1.9 SERT -4.7 -4.3 2.1 -7.6 -0.6 Opioid DOR -2.4 0.3 6.1 -0.2 0.6 KOR 23.7 26.8 27.2 30.0 7.0 MOR -1.8 -2.4 -2.1 1.3 1.5 GABA GabaA 27.4 21.7 9.6 16.1 18.1 Histamine H1 -5.7 -4.9 -4.8 -4.0 -5.1 H2 15.4 23.1 16.1 23.5 31.0 H3 10.1 10.4 11.5 5.1 16.2 H4 -10.7 4.3 -4.6 -3.6 -1.1 Muscarinic M1 -7.4 5.0 -5.3 -10.7 -2.2 M2 -18.2 -9.7 -12.4 -13.2 -14.1 M3 -16.2 -9.7 -11.9 -11.2 -12.2 M4 -14.6 -11.8 -16.5 -24.0 -14.7 M5 0.7 -5.8 -7.6 -9.1 9.3 Sigma Sigma1 33.6 23.0 30.8 25.8 30.9 Sigma2 -5.3 -5.7 -9.7 -8.5 -7.5

Secondary DAT binding assays of PGE2-EA and the azido compounds, AM7637 and

AM7638, were carried out to evaluate the potency of this observed interaction for the dopamine transporter. The resulting curves for AM7638, the 15R-hydroxy azido compound,

are shown in Figure 3.2. The secondary assay was completed twice; each time

administered alongside the reference compound, GBR12935, and an unrelated test

compound. The Ki was found to be 5.6 µM in the first assay and 4.6 µM in the second giving an average of 5.1 µM. Curves for PGE2-EA and AM7637 could not be completed as their

estimated Ki value exceeded 10 µM. Therefore, these compounds were not sufficiently

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potent at DAT to suggest meaningful activity or to give a basis to pursue these compounds as DAT inhibitors.

GBR12935 GBR12935 Ki=17nM Ki=10nM

AM7638 Ki=5.624µM AM7638 Ki=4.64µM

Figure 3.2 Radioligand binding curves for AM7638 (red in left; blue in right) and reference compound GBR12935 (black in both). (Other curve is unrelated test compound)

Alprenolol Ki=45nM

AM7638 Ki=5.624µM

Figure 3.3 Radioligand binding curve for AM7638 (red) and reference compound Alprenolol (black).

An additional secondary screen was successfully achieved by AM7638 on the beta3 adrenergic receptor. (Figure 3.3) Despite the primary screen giving only 16.5% inhibition at 10 µM, AM7638 exhibited a Ki value of 7.239 µM. The reason for the discrepancy is unknown.

Quantification of the affinity of AM7638 for the D5 dopamine receptor by secondary

screening was attempted, however, its estimated Ki was also determined to exceed 10 µM.

Additionally, all of our compounds were automatically subjected to secondary CB2 screening

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without an initial primary screen to establish activity. Not surprisingly, the Ki was greater

than 10 µM for all compounds. The results of the secondary screens are given in Table 3.2.

Table 3.2 Secondary radioligand binding assay results for Beta3, CB2, D5, and DAT receptors. Highlighted cells indicate radioligand inhibition potency (Ki) less than 10 µM. Given as Ki value (nM) (Radioligand competition) Receptor PGE2-EA AM7637 AM7638 AM7645 AM7646 Tested Final Azide S Final Azide R Final NCS R Final NCS S

Beta3 - - 7,239.0 - - CB2 >10,000 >10,000 >10,000 >10,000 >10,000 D5 - - >10,000 - - DAT >10,000 >10,000 5,132.5 - - (ave, n=2)

Finally, our compounds were screened in primary functional assays against cholecystokinin and several orphan receptors. These functional assays involve the monitoring of downstream GPCR signaling characteristic of Gs, Gq, or Gi/o to determine the

activity of compounds for the receptors. Unfortunately, the compounds showed no activity

towards any of the receptors. The test compounds’ percent efficacy or inhibition of the

activity shown by the receptors’ standard ligands did not exceed 0.7%. The results are

tabulated in Table 3.3.

Table 3.3 Primary functional assay results for cholecystokinin and 18 orphan receptors. Values are % of the efficacy shown by the known agonist/antagonist. Given as percent efficacy (agonist), percent inhibition (antagonist) Receptor Tested PGE2- AM7637 AM7638 AM7645 AM7646 EA Final Final Final Final Azide S Azide R NCS R NCS S CCK2 Agonist 0.1 - - - - CCK2 Antagonist 4.4 - - - - GPR1 0.3 0.3 0.3 0.3 0.3 GPR123 0.4 0.4 0.4 0.3 0.4 GPR132 0.4 0.3 0.2 0.3 0.3 GPR133 0.4 0.4 0.4 0.4 0.4 GPR157 0.4 0.3 0.3 0.2 0.3 GPR39 0.7 0.7 0.7 0.7 0.7 GPR41 0.2 0.1 0.1 0.1 0.2 GPR43 0.2 0.2 0.2 0.2 0.3 GPR45 0.2 0.3 0.2 0.2 0.3 GPR55 0.5 0.4 0.4 0.4 0.4 GPR57 0.7 0.7 0.7 0.7 0.7 GPR58 0.6 0.5 0.5 0.5 0.5

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GPR62 0.5 0.5 0.4 0.4 0.5 GPR68 0.3 0.3 0.3 0.3 0.4 GPR83 0.4 0.4 0.4 0.4 0.4 GPR84 0.3 0.3 0.2 0.3 0.3 GPR87 0.6 0.7 0.6 0.6 0.6 GPR88 0.2 0.2 0.2 0.3 0.3

From the results of these screens, we can conclude that none of the receptors

screened show particularly strong responses to our prostamide compounds. The observation that AM7638 exhibited low affinity for the DAT receptor introduces the intriguing possibility

that the endogenous prostamides are capable of inhibiting the synaptic uptake of dopamide.

However, as a singular target for the prostamide action, DAT does not represent a

promising lead. There exist many inhibitors of the dopamine transporter that are

substantially more potent. Interactions with orphan receptors would have solicited much more promising prospects, unfortunately, the prostamides were decidedly inactive against those orphan receptors that were screened. While the lack of hits found during the screening process is not surprising, the implementation of a full panel receptor screen is a fundamental starting place when exploring the activity of novel bioactive compounds.

All of the final prostamide products were screened against the FAAH enzyme for inhibition.† None of the compounds inhibited rat or human FAAH at concentrations up to 100

µM. Because the compounds did not compete with the fluorescent substrate, 7-amino-4- methyl coumarin-arachidonamide, for the active site, the conclusion was made that they did not bind FAAH. These results confirmed that prostamides were not substrates for FAAH and therefore showed enhanced stability towards hydrolysis to their prostaglandin counterparts.

Thus, any pharmacological effects attributed to treatment with prostamides are most likely due to prostamide’s biological actions and not due to prior conversion to prostaglandins.

† The FAAH screening was done by Erin Shelnut, Kyle Whitten, and Girija Rajarshi with assistance from Jodi Wood and Mahmoud Nasr

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3.2 Effects of Prostamide on Inflammation

The involvement of prostaglandins as pro-inflammatory, hyperalgesic lipid mediators

has long been recognized.1 However, more recent studies have established that

prostaglandins, specifically PGE2 and PGD2, trigger a class switch from pro-inflammatory to

pro-resolving lipid mediators.2,3 Counter to previous understanding, termination of inflammation is an active rather than passive process in which endogenous lipid mediators known as resolvins, protectins, and maresins act as anti-inflammatory and pro-resolving agents. The prostaglandins stimulate the transcriptional regulation of the 15-lipoxygenase pathway which, in turn, leads to greater concentration of the pro-resolving lipoxins and downstream resolvins. It is unknown whether the lipid mediator class switching initiated by

PGE2 and PGD2 is the result of interactions with the prostaglandin receptors themselves or

some other actions.

A clear indication of a system undergoing inflammation is the recruitment of polymorphonuclear neutrophils (PMN) associated with apoptosis.4,5 In order to return to homeostasis, PMN must be cleared from the affected tissue. The prostaglandins trigger the biosynthesis of the resolvins and protectins which in turn, diminish neutrophil infiltration and promote neutrophil clearance and release of anti-inflammatory and reparative cytokines like TGF-β1.6 An indication of a system under resolution of inflammation is the presence of

monocytes that aid in the clearance of PMN. Thus, monitoring PMN and monocyte levels in

tissues undergoing inflammation can give insight into whether pro-inflammatory or pro-

resolving processes are taking place. With this in mind, the next step in our study was to

observe the effects of the prostamide analogs within the inflammation model and compare

the results to those of PGE2.

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3.2.1 Reduction of polymorphonuclear neutrophils by prostamide treatment of zymosan A-induced inflammation

Figure 3.4 Compounds tested for inflammatory response

The actions of the 15S hydroxy, 20-isothiocyanato compound (AM7645) and PGE2 were assessed in vivo in a zymosan A-induced murine peritonitis model.† (Figure 3.4)

Peritonitis is an inflammation of the tissue lining the abdominal wall, and in this study, mice

received an intraperitoneal injection of the inflammation causing complex zymosan A. 5 minutes prior to the initiation of inflammation by zymosan A, 1 µg of either AM7645 or PGE2 was administered intravenously. After 4 hours, the standard amount time to allow for full inflammatory processes to take place, the mice were sacrificed and peritoneal exudates harvested. The neutrophils and monocytes were identified and enumerated by fluorescence- activated cell sorting (FACS) and light microscopy. Three mice were treated with PGE2, four were treated with AM7645, and four were left untreated as control models for inflammation.

† The in vivo mouse peritonitis studies were done by Dr Rong Yang in Serhan’s Lab at Harvard Med

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O H N OH O NCS HO OH AM7645

Control 1ug PGE2 1ug AM7645 n=4 n=3 n=4

Figure 3.5 Effects of PGE2 and AM7645 on cell numbers of PMN and monocytes injected with 1ug/mouse compound i.v., after 5min 1mg zymosan A. After 4hrs, leukocytes were identified/enumerated using light microscopy and fluorescence-activated cell sorting

The results of compound administration on cell numbers of (PMN) and monocytes are illustrated in Figure 3.5. Both PGE2 and AM7645 effectively reduce PMN infiltration in

murine peritonitis. Due to the knowledge that, in all cases, prostamides are less potent than prostaglandins at prostaglandin receptors, this result gives a powerful indication that the anti-inflammatory effects triggered by prostaglandin E2 and our prostamide compound are

not the result of activity at the prostaglandin receptor. Alternatively, it may suggest the

presence of two coexisting pathways recognizing prostaglandins and prostamides

separately. Interestingly, the results may indicate that AM7645 does not affect the number

of monocytes recruited into the tissue to assist in neutrophil clearance and resolution of

inflammation. PGE2, on the other hand, hinders monocyte levels to a small degree. Because

increased monocyte levels indicate an inflammation resolution process, unchanged levels or

increased levels are desirable for halting the inflammation process. We can conclude from

these studies that the prostamides are equally as effective anti-inflammatory, pro-resolving

agents as the prostaglandins have been demonstrated to be.

Intriguingly, because they are less potent at the prostaglandin receptors,

prostamides should not exhibit effects associated with EP receptor activation. The EP

receptors are generally associated with pain and inflammation. EP1 activation mediates

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inflammatory thermal hyperalgesia and neuroendocrine stress responses.7,8 EP2 facilitates

pain transmission in the central nervous system (CNS) and joint inflammation in collagen-

induced arthritis.9,10 EP3 activation leads to hyperalgesia and pyrogenic fever.11,12 And

finally, EP4 is also known to mediate joint inflammation in collagen-induced arthritis.10 Thus, due to the poor affinity for the prostamides for the prostaglandin receptors, they may carry the full benefit of prostaglandin’s ability to trigger a lipid mediator class switch from pro- inflammatory to anti-inflammatory without inducing the undesirable, pain-inducing side effects associated with prostaglandin receptor activation.

3.2.2 Determination of G protein signaling pathways by Electrical Cell-Substrate Impedance Sensing (ECIS)

To assess whether the effective reduction in PMN infiltration caused by the prostamide AM7645 is due to interactions with the prostaglandin receptors, in vitro screening assays were carried out using individual recombinant EP receptors in an electrical

cell-substrate impedance sensing system (ECIS).† ECIS is a cell based assay that can be used to investigate the signal transduction of G protein-coupled receptors such as the EP receptors.13 The assay was originally developed to monitor cellular growth and regrowth during wound healing. The electrical resistance or, in this case, impedance detected across the electrode surface of the circuit indicates the degree of coverage by growing cells.

(Figure 3.6) A larger impedance value denotes a greater number of cells are attached unto

the surface of the electrode.

† The ECIS studies were done by Dr Nan Chiang in Serhan’s Lab at Harvard Med

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Figure 3.6 (a) Schematic and image of an ECIS system 8-well plate and (b) a representation of the impedance caused by cellular cover of the detecting electrode.13

More recently it was discovered that ECIS assays can be used to study signaling

14-16 pathways of GPCRs. G proteins (Gs, Gi, or Gq) coupling to the receptors give diverse and characteristic profiles of impedance change upon ligand binding. Established patterns of impedance change traces have been defined and are demonstrated in Figure 3.7.13 GPCRs

with well-defined coupling exhibit marked impedance changes over time when treated with

their respective agonists. The Gs-coupled D5 dopamine receptor induces a substantial

decrease in cellular impedance when treated with dopamine. This response is typical of all

Gs-coupled GPCRs. The D2 dopamine receptor, alternatively is a well-known Gi/o-coupled

receptor and when provided dopamine ligand yields a clear increase in impedance characteristic of all Gi/o receptors. The remaining GPCR type, Gq, is represented by the M1 muscarinic receptor which elicits a signature decrease in impedance followed by an overall increase when treated with acetylcholine. The mechanism by which G protein signaling affects impedance is not well understood, however alterations of the cellular cytoskeletal elements are evidently responsible. Preliminary studies show that changes in actin dynamics responsible for cell junctions and shape may be involved in the variations in impedance.17-19

Regardless of the mechanism, with these patterns established compounds can be tested against GPCRs and their downstream signaling effects can be directly measured.

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a)

Gs D5, dopamine ↑ cAMP ↑ adenylate cyclase

b)

Gi/o D2, dopamine ↓ cAMP ↓ adenylate cyclase

c)

Gq M1, ACh ↑ phospholipase C (PLC) ↑ inositol triphosphate

(IP3)

Figure 3.7 Cellular impedance profiles unique to each of the G signaling pathways. (a) Gs impedance profile is represented by dopamine treatment of D5 in CHO cells and gives a signature decrease in impedance. (b) Gi/o impedance profile is represented by dopamine treatment of D2 in CHO cells and gives a characteristic increase in impedance. (c) Gq impedance profile is represented by acetylcholine treatment of M1 in CHO cells and gives an initial decrease in impedance followed by a marked increase.13

ECIS gives a unique advantage over standard binding assays in that the functionality of the agonist activity is measured and not simply the binding ability. Whether or not a compound binds well to a receptor does not dictate its overall cellular effects. By directly evaluating the downstream functional signaling, the compounds that bind the same receptor but elicit different responses can be distinguished. This is very informative in our case, as prostamides and prostaglandins have both been shown to have some ability to bind

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prostaglandin receptors, however their observed pharmacology, as discussed in the

background section, is diverse.

A complication to the study of GPCRs is their pleiotropic nature. Depending on the

cellular environment in which the receptors are resident, a single agonist can give different

G protein responses. Receptors expressed in HEK cells may give Gs-coupled impedance

profiles, while those same receptors expressed in CHO cells give a Gi/o-coupled profile of impedance. This phenomenon is extended to endogenous cells. In addition, identical

Gs D5, dopamine ↑ cAMP ↑ adenylate cyclase

Gi/o D2, dopamine ↓ cAMP ↓ adenylate cyclase

Gq M1, ACh ↑ phospholipase C (PLC) ↑ inositol triphosphate

(IP3)

Figure 3.8 Effects of pertussis toxin (PTX) and Cytochalasin D on the cellular impedance profiles unique to each of the G signaling pathways. PTX reverses only the Gi/o impedance change. Cytochalasin cancels out the Gs and the Gi/o impedance change but only slightly hinders the Gq impedance change.13

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receptors expressed within the same cell type may give various downstream signaling pathways when treated with different agonist. In fact, to further complicate matters, some receptors exhibit activation of multiple pathways when agonized. Fortunately, the impedance profiles of these receptors are recognizable from their shape and can be deduced simply by treating with direct inhibitors of one of the G-proteins. (Figure 3.8) Gi/o responses can be negated by treatment with pertussis toxin (PTX), and Gq impedance is the

only trace left unchanged by treatment with cytochalasin D. From these distinguishable traits, GPCRs that exhibit a combination of signaling pathways can be inferred. For example, the CB1 receptor is known to exhibit both Gs and Gi/o patterns in its impedance trace.

(Figure 3.9) Depending on its expression in either HEK or CHO cells, two different patterns

emerge. In HEK cells there is an initial marked decrease in impedance followed by a steady

increase back towards baseline. CB1 receptors expressed in CHO cells, on the other hand, have an overall moderate and slow increase in impedance. These traces are reflective of a combination Gs-Gi/o-coupled signaling pathway. When treated with PTX, the Gi/o contributing

signal is blocked and only the Gs trace remains.

a b

Figure 3.9 Novel impedance profiles of the CB1 receptor treated with CP55940 and Win55212-2

agonists and expressed in (a) CHO cells and (b) HEK cells due to competing Gs and Gi/o signaling

pathways. Treatment with PTX reverses the Gi/o response leaving only the impedance profile of Gs signaling.13

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3.2.3 Implementation of ECIS to Determine the Activity of Prostamide on the Prostaglandin Receptors

In the present case, ECIS was used to qualitatively compare the G-protein response

of the prostaglandin receptors activated by PGE2 to that induced by the prostamide test compound, AM7645. Employing a single receptor in a single cell type should allow us to compare the agonist responses of the two compounds. If the impedance caused by AM7645 resembles that of PGE2, it can be assumed that both are active at the given receptor and elicit the same downstream signaling pathways. If the impedance trace is dissimilar to the

PGE2 but follows a pattern comparable to one of the other characteristic G-protein impedance traces, the compound is in fact acting on the receptor but producing unique signaling. Finally, the possibility arises that AM7645 does not activate the prostaglandin receptors and in this case the impedance will resemble treatment with vehicle giving a baseline response.

The results confirm prostamides are inactive at the EP prostaglandin receptors, and thus their elicited effects on PMN infiltrate are not due to stimulation of these receptors.

(Figure 3.10) This lack of prostaglandin E receptor activation by our prostamide is

20-23 consistent with studies previously discussed in Section 1.4.1.1. PGE2 gives a marked

24 Gq-like change of impedance trace characteristic of EP1 activation. (Figure 3.10a)

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a b

c d

e f

Figure 3.10 Ligand-receptor dependent changes in impedance monitored using an Electrical Cell- Substrate Impedance Sensing System (ECIS). CHO cells transfected with EP1 (a,b), EP3 (c,d), or EP4 (e,f) receptors and plated unto an 8-well ECIS chamber (8W10E+) at 0.2x106 cells/well were treated with 100nM PGE2 (a,c,d) or with PGE2-EA or AM7645 (b,d,f).

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Conversely, PGE2-EA and AM7645 show very muted changes in impedance similar to that of vehicle. (Figure 3.10b) Likewise, PGE2 demonstrates the ability to activate EP3 providing a

distinctive Gi/o-like ECIS trace, whereas the prostamides give a trace nearly identical to

24 vehicle. (Figure 3.10c,d) Finally, when EP4 was treated with PGE2, the appropriate Gs-like

24 activity resulted. When treated with PGE2-EA and AM7645 these effects were absent.

(Figure 3.10e,f)

In conclusion, the prostamides have shown the ability to influence inflammation by

enhancing the clearance of polymorphonuclear neutrophils that is required for resolution of

the inflammatory state. However, according to ECIS studies these effects are likely not the

result of activity at the prostaglandin receptors.

3.3 Effects of Prostamide on Immunosuppression

One of the first responses of the immune system to injury or infection is

inflammation. The AM7645 displayed the potent ability to mediate neutrophil infiltration

without affecting pro-resolving monocyte levels. To explore the possibility that the

prostamides may have further immunomodulatory effects, our study was extended to

monitor the impact of our compounds on lymphocyte production.† The key cellular barrier in immunological response, lymphocytes also known was white blood cells, include natural killer cells (NK), thymus cells (T-cells), and bursa-derived cells (B-cells) and act to identify

and eliminate pathogens. Compounds eliciting an increase in the production of lymphocytes

support the immune system response and those that decrease levels are described as

immunosuppressive.

In order to determine the influence of the prostamides on lymphocyte production,

our compounds AM7637, AM7638, and AM7645 and PGE2-EA were tested in two distinct in

vitro assays. (Figure 3.11) The first to be carried out was a plaque-forming cell assay

† The leukocyte studies were done by Joseph Meissler in Eisenstein’s Lab at Temple University

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Figure 3.11 Compounds tested for immunosuppression

(PFC) in which mouse spleen cells are incubated with sheep red blood cells (SRBC) in cell

culture for five days. The SRBCs serve as antigens to the lymphocytes produced by the

mouse spleen cells. The lymphocytes that respond to SRBCs differentiate and secrete

antibodies. Quantification of these lymphocytes is achieved by harvesting the cells, adding

to them an excess of SRBCs plus guinea pig complement, and distributing between two

glass slides. The complement attaches to the antibodies and allows for mediated lysis of the antibody-coated SRBCs resulting in the formation of plaques in the layer of cells. The plaque is formed by lysed SRBCs surrounding a single antibody-secreting lymphocyte. Thus, the antibody-secreting lymphocytes can be identified by the surrounding plaques and easily enumerated under dissecting microscope.

Upon treatment with either vehicle or various doses of the prostamide test compounds, the effects on the number of plaques resulting from the antibody-secreting lymphocytes can be studied. A decrease in the number plaques formed would indicate a reduction in the amount of lymphocytes present and would represent immunosuppression.

Alternatively, an increase in plaques would signify an enhancement of immune response.

The results are expressed as a suppression index (SI) which equates to the number of plaque-forming cells when treated with test compound divided by the number of plaque- forming cells without treatment (control). A value of one would signify no change in the

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immune response, a value below one would mean immunosuppression, and a value above one would mean augmentation.

The results shown in Figure 3.12 reveal that all of the prostamides tested are substantially immunosuppressive. The 15R,20-azido compound (AM7638) gave the greatest reduction in plaque formation down to 10 µM, and the 15S,20-azido compound (AM7637)

was also quite robust at 10 µM. Meanwhile, the 20-isothiocyanato compound (AM7645) and

the standard prostamide E2 (AM7647) were effective only at concentrations above 24 µM.

Thus, the azido compounds exhibit a more potent immunosuppressive effect than the isothiocyanato and standard prostamide compounds. The 15R isomer was slightly more

potent in evoking immunosuppression than the 15S isomer, however whether this result

represents a substantial distinction remains to be seen. DMSO vehicle also exhibited

immunosuppressive behavior, but not nearly to the extent of the prostamides.

Figure 3.12 Dose response curves of AM7638, AM7645, AM7637, PGE2-EA (AM7647) in a plaque- forming cell assay (PFC) quantifying the ratio of the number of plaques formed by lymphocytes when treated with the test compounds compared to the number of plaques formed without treatment.

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PFC is a valuable method of determining the overall effect of a given test compound on general immune response. Effective plaque formation requires functional antigen presenting cells such as macrophages and dendritic cells and lymphocytes such as T-cells and B-cells. Therefore, the resulting change in plaque formation may be due to prostamide’s influence on any or all of the cells involved.

The second study completed to determine prostamide’s effect on immunosuppression

was a mixed lymphocyte reaction assay (MLR) which focuses mainly on the suppression of

T-cells. A common method used to assess immune response, MLR involves the mixing of

lymphocyte cell cultures from two individual histoincompatible mouse strains. Due to the

incompatibility, the T-cells recognize those cells from the other strain as foreign and in

response are activated to proliferate. The assay monitors the amount of proliferation by

measuring the uptake of tritiated thymidine. Thymidine is a nucleoside of DNA and is

essential to the dividing of cells during proliferation. Thus, when the system is treated with

an immunosuppressive compound, less proliferation is evident by the reduced uptake of

tritiated thymidine during MLR. A compound that is determined to support the immune

system would increase proliferation and the observed uptake of thymidine.

The MLR study results revealed that our prostamide compounds had much less of an

effect on immunosuppression than that determined by PFC. Of all the compounds tested,

AM7637, the 15S, 20-azido analog, exhibited the most potent immunosuppressive behavior.

(Figure 3.13) Giving approximately 30% inhibition of the lymphocyte proliferation, in a

range of 0.1-1 µM. Tetrahydrocannabinol (THC) serves as a positive reference compound

that, similar to CB2 selective agonists, exhibits maximum immunosuppression between 10

and 32 µM. Because test compound concentrations above 10 µM were not used, it is

possible that these prostamide might still affect immunosuppression with potency similar to

the CB2 agonists. Based on these assays, AM7645 and PGE2-EA both have the ability, at

higher concentrations, to substantially suppress the lymphocyte immune response.

Interestingly, the 15R azido analog showed the least amount of change to the immune

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AM7647 = PGE2-EA

AM7647 = PGE2-EA

Figure 3.13 Dose response curves of AM7638, AM7645, AM7637, PGE2-EA (AM7647), and reference compound THC in a mixed lymphocyte reaction assay (MLR) monitoring tritiated thymidine levels.

response, suggesting perhaps that the orientation of the 15-hydroxyl may, in fact, be a key determinant of immunomodulation.

From this research it can be concluded that prostamides have the ability to suppress the immune system although not to the same extent as the CB2 agonists such as THC. Both the PFC and MLR assays demonstrated that the prostamides limit the production of lymphocytes that are required for immunological response. Greater activity was observed in the PFC assay perhaps due to the additional target cells such as macrophages and B-cells within this test.

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3.4 Summary of Results

The receptor screening results revealed that the prostamide analogs tested are

inactive at almost all of receptors screened including the adrenergic, cannabinoid,

dopamine, GABA, histamine, muscarinic, serotonin, sigma, opioid, cholecystokinin,

norepinephrine transporter, and serotonin transporter receptors. However, there were two

exceptions as the prostamides showed some enhanced affinity for the dopamine transporter

receptor, DAT, and the beta3 adrenergic receptor. DAT is a membrane-spanning protein that facilitates the reuptake of dopamine from the synapse back into the cytosol for

degradation.25,26 It is the primary mechanism through which dopamine is cleared from the

synapses. Inhibition of this transporter by prostamide may increase synaptic levels of

dopamine and create a euphoric effect commonly associated with DAT inhibitors. This is an

interesting observation in that endogenous prostamide may indeed act to inhibit dopamine

reuptake, however this is most likely not prostamide’s primary biological purpose.

Conversely, the beta3 adrenergic receptor is known to facilitate lipolysis in adipose cells and

27 selective agonism of beta3 has been postulated as a treatment for obesity. Inhibition of the beta3 receptor by prostamide may decrease lipolysis in adipose cells and facilitate

weight gain. The inability of the prostamide analogs to inhibit FAAH and MGL enzymes confirms that prostamides are not substrates for metabolism by these enzymes and supports the assertion that prostamides exhibit substantial metabolic stability.

The results of prostamide analog treatment during inflammatory challenge indicated that prostamides are equally effective anti-inflammatory, pro-resolving agents as the prostaglandins. However, it is unlikely that the pro-resolving effects elicited by the prostamides are the result of activity at the prostaglandin receptors. Prostamides are known to be substantially less potent than prostaglandins at prostaglandin receptors, thus if prostaglandin receptor activation were required for resolution, one would expect prostamides to exhibit greatly reduced efficacy in decreasing PMN infiltration. This was not

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the case, however as both prostaglandin and prostamide enhanced PMN infiltration to

similar extents. In fact, results from ECIS assays confirmed that the prostamide analog tested was entirely inactive at the EP prostaglandin receptors. These results are especially intriguing as they may indicate the prostamides bring the full benefit of prostaglandin’s ability to trigger lipid mediator class switching from pro-inflammatory to pro-resolving without inducing the undesirable nociceptive side effects associated with prostaglandin receptor activation.

Finally, the immunoassay results suggest that the prostamides exhibit a mild suppression of the immune system. Lymphocyte production was reduced by treatment with the prostamide analogs in both the PFC and MLR assays.

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2. Levy, B. D. C., C. B.; Schmidt, B.; Gronert, K.; Serhan, C. N. . Nat. Immunol. 2001, 2, 612.

3. Bandeira-Melo, C.; Serra, M. F.; Diaz, B. L.; Cordeiro, R. S. B.; Silva, P. M. R.; Lenzi, H. L.; Bakhle, Y. S.; Serhan, C. N.; Martins, M. A., Cyclooxygenase-2-derived prostaglandin E2 and lipoxin A4 accelerate resolution of allergic edema in Angiostrongylus costaricensis- infected rats: relationship with concurrent eosinophilia. J. Immunol. 2000, 164, 1029.

4. Segal, A. W., How neutrophils kill microbes. Annu. Rev. Immunol. 2005, 23, 197.

5. Underhill, D. M.; Ozinsky, A., Phagocytosis of microbes: complexity in action. Annu. Rev. Immunol. 2002, 20, 825.

6. Levy, B. D.; Clish, C. B.; Schmidt, B.; Gronert, K.; Serhan, C. N., Lipid mediator class switching during acute inflammation: signals in resolution. Nat. Immunol. 2001, 2, 612.

7. Moriyama, T.; Higashi, T.; Togashi, K.; Iida, T.; Segi, E.; Sugimoto, Y.; Tominaga, T.; Narumiya, S.; Tominaga, M., Sensitization of TRPV1 by EP1 and IP reveals peripheral nociceptive mechanism of prostaglandins. Mol Pain 2005, 1, 3.

8. Matsuoka, Y.; Furuyashiki, T.; Bito, H.; Ushikubi, F.; Tanaka, Y.; Kobayashi, T.; Muro, S.; Satoh, N.; Kayahara, T.; Higashi, M.; Mizoguchi, A.; Shichi, H.; Fukuda, Y.; Nakao, K.; Narumiya, S., Impaired adrenocorticotropic hormone response to bacterial endotoxin in mice deficient in prostaglandin E receptor EP1 and EP3 subtypes. Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 4132.

9. Reinold, H.; Ahmadi, S.; Depner, U. B.; Layh, B.; Heindl, C.; Hamza, M.; Pahl, A.; Brune, K.; Narumiya, S.; Mueller, U.; Zeilhofer, H. U., Spinal inflammatory hyperalgesia is mediated by prostaglandin E receptors of the EP2 subtype. J. Clin. Invest. 2005, 115, 673.

10. Honda, T.; Segi-Nishida, E.; Miyachi, Y.; Narumiya, S., Prostacyclin-IP signaling and prostaglandin E2-EP2/EP4 signaling both mediate joint inflammation in mouse collagen- induced arthritis. J. Exp. Med. 2006, 203, 325.

11. Ueno, A.; Matsumoto, H.; Naraba, H.; Ikeda, Y.; Ushikubi, F.; Matsuoka, T.; Narumiya, S.; Sugimoto, Y.; Ichikawa, A.; Oh-ishi, S., Major roles of prostanoid receptors IP and EP3 in endotoxin-induced enhancement of pain perception. Biochem. Pharmacol. 2001, 62, 157.

12. Ushikubi, F.; Segi, E.; Sugimoto, Y.; Murata, T.; Matsuoka, T.; Kobayashi, T.; Hizaki, H.; Tuboi, K.; Katsuyama, M.; Ichikawa, A.; Tanaka, T.; Yoshida, N.; Narumiya, S., Impaired febrile response in mice lacking the prostaglandin E receptor subtype EP3. Nature (London) 1998, 395, 281.

13. Peters, M. F.; Scott, C. W., Evaluating cellular impedance assays for detection of GPCR pleiotropic signaling and functional selectivity. J. Biomol. Screening 2009, 14, 246.

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21. Woodward, D. F., Pharmacological Characterization of a Novel Antiglaucoma Agent, Bimatoprost (AGN 192024). J. Pharmacol. Exp. Ther. 2003, 305, 772.

22. Glass, M., Misidentification of prostamides as prostaglandins. J. Lipid Res. 2005, 46, 1364.

23. Ross, R. A., Pharmacological Characterization of the Anandamide Cyclooxygenase Metabolite: Prostaglandin E2 Ethanolamide. J. Pharmacol. Exp. Ther. 2002, 301, 900.

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26. Norregaard, L.; Gether, U., The monoamine neurotransmitter transporters: structure, conformational changes and molecular gating. Curr. Opin. Drug Discovery Dev. 2001, 4, 591.

27. Ferrer-Lorente, R.; Cabot, C.; Fernandez-Lopez, J.-A.; Alemany, M., Combined effects of oleoyl-estrone and a β3-adrenergic agonist (CL316,243) on lipid stores of diet-induced overweight male Wistar rats. Life Sci. 2005, 77, 2051.

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Chapter 4: Future Directions for Prostamide Research

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The studies presented within this dissertation represent the earliest stages of investigation into prostamide pharmacology. A considerable amount of work remains to further elucidate the physiological function of the prostamides and to advance our understanding of the therapeutic possibilities of prostamide compounds. Areas of study in which the knowledge of prostamide activity can be further expanded include neuroprotection and suppression of production in the central nervous system,1-3 activation

of individual EP receptors in isolated functional tissue assays,4 recognition of prostamide by

EP splice variant proteins,5 facilitation of resolution processes in inflammation models,

suppression of the immune system response, recognition of prostamide by nuclear receptors

6,7 such as PPARγ, and isolation of prostamide-selective receptors from prostamide-sensitive

tissue.

Several physiological responses have been shown to result from treatment with

prostamides including protection of neurons of the central nervous system (CNS),1-3 reduction of intraocular pressure in glaucoma,8,9 facilitation of polymorphonuclear neutrophil

clearance during inflammation (Section 3.2.1), and suppression of immune response

(Section 3.3). Submitting our modified covalent probes to further testing within these systems and comparing their effects to those of prostaglandins may lead to key discoveries

in the unique actions of the prostamides. Tissues that show an enhanced sensitivity to

activation by prostamides such as feline lung parenchyma,8 rabbit uterus,10 and feline

iris.9,11 can be used in the identification and isolation of putative prostamide-selective target

proteins.

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4.1 General Methods for Isolating Prostamide-Selective Target Proteins from Prostamide-Sensitive Tissue

4.1.1 Binding Specificity Assay

A general method for the detection of receptors within a tissue that responds to prostamide entails a membrane preparation first being subjected to a specific binding assay

using tritiated prostamide E2 and screening the covalent prostamide probes for inhibitory activity.12 The prostamide-selective tissue chosen to provide the membrane preparation serves as the source of the putative prostamide receptors. The probes’ ability to compete

3 with tritiated prostamide E2 ([ H]PGE2-EA) upon incubation, and thus act an inhibitor within

the tissue, gives a direct measure of the probe’s potency. An example of feasible curves

3 resulting from a [ H]PGE2-EA binding specificity assay is illustrated in Figure 4.1.

Prostamide E2 itself would likely be the most potent inhibitor of the tritiated prostamide E2

radioligand. With binding curves in hand, the affinity of our probes for the putative

prostamide receptors can be ascertained. Further design of analogs based on the SAR may

lead to the discovery of a probe with substantially enhanced affinity for the prostamide

receptor.

Figure 4.1 A representation of curves that may result from a membrane specific binding assay of prostamide analogs on putative prostamide receptors. PGE2-EA will most likely exhibit the highest potency.

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While the tissues selected for these specific binding assays are those outlined above

that exhibit enhanced sensitivity towards prostamide, it is expected that some of the

activity achieved within this assay may be due to interactions with prostaglandin receptors

within the same tissue. To estimate the magnitude of these offsite interactions, standard

prostaglandin compounds such as PGE2 and 17-phenyl PGE2 should also be screened for specific binding. If their ability to inhibit tritiated prostamide E2 binding is similar or more

potent than that of PGE2-EA, the specific binding curves for the prostamides are most likely

the results of the probes binding mainly to prostaglandin receptors. However, if

prostaglandins potency is substantially lower than that of the prostamides the evidence would suggest that the tissue does in fact contain proteins that selectively recognize prostamide over prostaglandins. Because offsite prostamide activity at prostaglandin receptors complicates the observed activity within the specific binding assays, the IC50 values determined by the curves should not be taken as the accurate potency of the

compound for the putative prostamide receptor alone but instead mainly as an indication of

the presence of proteins within the tissue that selectively recognize the prostamides.

To address the possibility that the prostamides are acting solely at prostaglandin

receptors within the given tissue, the effects of administering individual EP receptor

antagonists along with prostamide test compounds in the given tissue response would

indicate their involvement in prostamide activity. Several selective antagonists have been

developed for each of the four EP receptor subtypes. ONO8711 and SC51089 antagonize

13,14 15,16 17,18 EP1, AH6809 and PF04418948 antagonize EP2, DG041 and CM9 antagonize EP3,

19,20 and AH23848B and CJ023423 antagonize EP4, all selectively. (Figure 4.2) By

antagonizing each receptor individually within the given tissue and determining their ability

to block the response incited by prostamide E2 and the prostamide probes, their distinct participation in prostamide signaling can be resolved. The degree to which the response is antagonized indicates the amount of prostaglandin receptor involvement. If the antagonists

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Figure 4.2 Structures of the selective EP prostaglandin receptor antagonists

do not entirely block prostamide activity, there is evidence that there may be a separate

target with which prostamide is interacting. However, if antagonism of one or more EP subtype blocks nearly all prostamide activity, prostamide is most likely eliciting its effects within that tissue by acting exclusively on prostaglandin receptors.

4.1.2 Bifunctional Prostamide Probes

4.1.2.1 Isolation of Prostamide-Selective Proteins using Radiolabeled Probes

Assuming one or more of the covalent probes achieves sufficient activity within the specific binding screens and the EP receptor antagonists do not entirely block prostamide

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activity, the next step in the isolation of the prostamide selective receptor is radiolabeling

the targeted proteins. Bifunctional probes containing both a covalent attachment modification and a radiolabel such as 125I can serve to essentially tag the proteins covalently modified by the ligand for effective purification.12 (Figure 4.3) 125I is the preferred radioisotope for protein labeling experiments due to its substantially long half-life of 60

days. A proposed synthetic route to the radioiodination of this probe is outlined in Section

4.8.2.1. Radiolabelling allows for identification of the proteins to which the ligand binds

following separation of the proteins by polyacrylamide gel purification (PAGE)

electrophoresis. Subsequent to isolation of the radioactive PAGE bands, characterization of

the prostamide receptor proteins can be performed using various mass spectrometry

techniques. Essential information including the proteins’ overall molecular weight, the

identity of the amino acid to which the covalent ligand binds, and the amino acid sequence

can be obtained from these studies. Comparison of the results to those of the well-

established prostaglandin receptors may lead to the discovery of novel receptors targeted

by the prostamides.

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Figure 4.3 Application of bifunctional covalent, radiolabeled probes for isolation and identification of prostamide-binding proteins. Following successful covalent attachment of the probe, gel purification of the proteins leads to numerous bands. Autoradiography allows for the identification of the bands resulting from radioactively bound protein. Scintillation spectroscopy gives a quantifiable radioactivity to each fraction. Upon separation, the protein can undergo mass spectrometry to aid in determining its identity. Adapted from Kawada, 1989.12

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4.1.2.2 Emission Topography Imaging using Radiolabeled Probes

Figure 4.4 Examples of radioactive tomography imaging of human brain. (A) PET image of the distribution of D2 dopamine receptors in a normal human brain using [11C]-FLB 457;21 (B) SPECT image of the distribution of the benzodiazepine site of the GABAA receptor in a human brain 2 months 123 22 after trauma using [ I]-iomazenil; (C) autoradiography image of the GABAB receptor distribution in a section of post-mortem human brain using [3H]-CGP 54626.23 Colors represent amount of binding, with red as high and blue as low.

In addition, treatment of radioiodinated probes in vivo allows for positron emission

tomography (PET) and single photon emission computed tomography (SPECT) imaging.24-27

Both of these imaging techniques allow for study of the biodistribution of intravenously administered radioligand and are particularly useful in neuroendocrine applications. Three dimensional images are rendered based on a series of scans that measure gamma radiation levels. (Figure 4.4) In this way, the location of increased concentrations of the

radiolabelled prostamides can be ascertained and potential cell populations that show

enhanced interactions with the probes can be identified. Information pertaining to the

prostamide’s pharmacokinetics can also be monitored through tomography imaging. The principal differences between the two imaging techniques are the radioisotopes used and the resolution of the rendered image. In addition to 124I which exhibits a relatively long half-

life of four days, PET imaging commonly employs 11C and 18F with half-lives of twenty minutes and about two hours, respectively. Conversely, SPECT imaging utilizes 131I as its major radioisotope which has a half-life of about eight days. This substantially long half-life

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is advantageous due to increase in time available for synthesis, purification, and

administration of the radiolabeled probes, however SPECT tends to give images that have a

lower resolution than those of PET imaging. Autoradiography can be used to produce a two

dimensional image of the radioisotope concentrations in slices of isolated tissues.

4.1.2.3 Affinity Purification of Prostamide-Selective Proteins using Biotinylated Probes

Alternatively, a diverse series of bifunctional probes could be designed incorporating biotin labels in place of radiolabels. The biotin label is commonly employed in the isolation of proteins due to their ease of introduction, small size, and high affinity and specificity for the

proteins avidin or streptavidin.28,29 (Figure 4.5) A suggested synthetic route to the biotinylation of this probe is described in Section 4.8.2.2. In this type of affinity purification, the covalent probe essentially fastens the biotin label to the recognizing protein, and the linked biotin is then able to bind avidin protein coated onto resin beads.30

Thus the avidin beads capture only the proteins that effectively bind prostamide, and all other unbound proteins are washed away. (Figure 4.6) The biotin/avidin complex can be cleaved at elevated temperatures using dithiothreitol (DTT). The released biotinylated ligand-bound protein can then undergo separation in PAGE by electrophoresis.

Figure 4.5 The chemical structure of biotin and the three dimensional crystal structure of its target protein, avidin. Biotin binds avidin with extremely high affinity and selectivity, and four molecules of biotin can bind each avidin protein.

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Figure 4.6 (A) Representation of a bifunctional biotin-labeled covalent prostamide probe; (B) Application of biotin-labeled probes for isolation and identification of prostamide-binding proteins. Following successful covalent attachment of the probe, the biotin-labeled protein is introduced to an avidin-coated resin bead. The beads are washed to remove any non-bound proteins, and thus only proteins that recognize the prostamide probes remain. Gel purification and in-gel digestion of the isolated bands gives samples for mass spectrometry. The protein segments resulting from the digestion can be studied by mass spectrometry to identify the overall sequence and identity of the proteins.

4.2 Prostamide E2 Activity in the Central Nervous System

One of the most promising areas in which prostamide’s physiological effects may lead to identification of key targets is within the CNS where the concentration of prostamide’s precursor anandamide is high and where recent studies have shown prostamides to enhance neuroprotection and neuroplasticity.1-3 While the endocannabinoids, anandamide and 2-arachidonoylglycerol, have long been attributed the ability to protect neuronal cells, these recent studies have shown that the neuroprotective effects are in fact the result of anandamide’s metabolic product prostamide and not anandamide itself.

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4.2.1 Neuroprotection by Prostamide E2

Further study into the neuronal effects of the prostamides using our covalent probes

may lead to the source of prostamide’s protective capabilities. Using conditions similar to

those of the study detailed in Section 1.6.3 by Bobrov’s group at the Russian Academy of

Medical Sciences in Moscow, neurons undergoing reduced K+-triggered apoptosis are treated with the prostamide probes and the cell survival quantitated.2 If neuronal survival increases with prostamide treatment, as we would expect based on the literature, the

probes are effectively enhancing the same neuroprotective pathways. The ability of the

probes to covalently bind the proteins may increase the resulting neuroprotection over time

and additionally, allow for isolation of the target proteins responsible for the protective

effects. Within this study, increase or decrease in neuron survival can be used to screen a

small set of prostamide analogs in an SAR-like manner. The probes that most potently

increase cell survival can be used in further studies incorporating bifunctional moieties to aid

in isolation of the target proteins.

To determine if the neuroprotective properties are unique to the prostamides,

prostaglandin compounds such as PGE2 and 17-phenyl PGE2 should be tested along with the probes. Additionally, studying the administration of arachidonic acid and anandamide in the

presence and absence of a COX inhibitor may shed more light on which class of lipids exhibit

the greatest neuroprotective outcome. Similar enhancement of neuron survival by the

prostaglandins to that of the prostamides may mean that there is an overlapping of activity

between the two sets of eicosanoids. However, if prostaglandins display less potent or less

efficacious protection of the neurons, this may constitute a key discovery in the unique

physiological function of the prostamides.

Finally, monitoring neuron survival upon administration of the individual EP

antagonists and treatment with prostamide E2 gives a clear indication of whether action at

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the prostaglandin receptors alone is likely responsible for the neuroprotective properties of

prostamide. Employing antagonists (Figure 4.2) for each EP receptor subtype separately

and quantifying neuron cell survival indicates any portion of prostamide’s activity attributed

to interaction at the prostaglandin receptor. If antagonism of a single receptor subtype or a

combination of two or more receptor subtypes can block all neuroprotection elicited by

prostamide, the neuroprotective effects of prostamide can be attributed entirely to activation of the prostaglandin receptors. If this were to be the case, prostaglandin E2 and

17-phenyl prostaglandin E2 would be expected to provide exceptional neuroprotection. If however, the prostamide probes exhibit residual neuroprotective activity following treatment with the prostaglandin receptors, the evidence would suggest a presence of prostamide selective targets.

Isolation of the target proteins within the nervous tissue requires the use of bifunctional probes as introduced in Section 4.1.2. Assuming the prostamide probes demonstrate effective increases in neuron cell survival, they can be tested for their ability to covalently bind the target proteins. When sufficient covalent modification is achieved, the proteins within the nervous tissue, to which the prostamides bind, can be “fished out” using either radiolabelled or biotinylated bifunctional covalent prostamide probes.

4.2.2 Interleukin-12 Suppression by Prostamide E2

A second study in favor of the neuroprotective properties of prostamide over those of anandamide led to the discovery that prostamide E2 is essential for the suppression of the

IL-12 cytokine. Using methods similar to those of Guaza’s group at the Cajal Institute in

Madrid described in Section 1.6.3, the influence of the prostamide probes on IL-12

production in macrophages and microglial cells can be determined by monitoring p40, p35,

and p19 subunit levels.1 The quantification of IL-12 subunit levels can be used as an assay

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against which the prostamide analogs can be screened. Probes that potently inhibit IL-12

production can again be adapted to bifunctional probes and used in isolating potential

receptor proteins from the macrophages and microglial cells.

The literature suggests that the EP2 receptor, and not the EP4 receptor, is involved in

the inhibitory actions of prostamide E2 due to the ability of AH6809, and not AH23848B, to

1 partially reverse prostamide’s reduction of IL-12. Thus, EP2 has been identified as part of the pathway through which prostamide down-regulates IL-12. Further studies can be done to determine whether EP1 and EP3 also play a role in the regulation of IL-12 by administering selective antagonists for these receptors. Additionally, prostaglandin E2 itself

should be tested for inhibition of IL-12 production and the results compared against those of

prostamide E2. If prostaglandin receptors constitute the extent of prostamide’s interactions within this system, prostaglandin analogs should have an enhanced ability to inhibit IL-12.

In this case, prostamides would serve strictly as prostaglandin-mimetics and their advantage over the prostaglandins themselves, within the CNS, would only be derived from

their superior availability and metabolic stability.

4.2.3 Interleukin-2 Suppression by Prostamide E2

In addition to inhibiting IL-12, prostamides also potently suppress the proliferation of

T-cells by inhibiting IL-2 production in murine splenocytes. Following methods similar to those used by Kaminski’s group at Michigan State University in East Lansing, Michigan, and discussed in Section 1.5.2, an enzyme-linked immunosorbent assay (ELISA) can be used

to monitor IL-2 in monocytes and macrophages upon treatment with the prostamide

probes.6 The interactions with prostaglandin receptors can be determined by co-treatment with the prostamide analogs and prostamide selective antagonists. The most potent analogs

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can then be converted to bifunctional radiolabelled or biotinylated probes and used to

isolate receptors within the monocyte and macrophage cells.

4.3 Prostamide Activity in Functional Tissue Assays

Functional tissue assays similar to those completed in Ross’s group at the University of Aberdeen in Scotland can be revisited employing our probes and the prostamide receptor antagonists.4 Ross’s original study, discussed in Section 1.4.1.1, establishes the overall

enhanced activity of the prostamides in tissues expressing the EP receptors compared to the

isolated EP receptors. Treating each of these tissues with prostamides in the presence and

absence of the prostaglandin receptor antagonists will lead to a better understanding of the

distinction between prostamide and prostaglandin action. If the contraction or relaxation response is completely inhibited by the prostaglandin antagonist, the prostamide is acting entirely at the prostaglandin receptors. If however, when prostamide probes and

prostaglandin receptor antagonists are co-administered there is residual activity, it is likely

that there is a separate set of proteins with which the prostamides are interacting. Thus

studies can be attempted using guinea pig trachea, known to express EP1 and EP2, guinea

pig vas deferens, known to express EP3, and rabbit jugular vein, known to express EP4. The

contraction and relaxation responses incited by treatment with prostamide are monitored

with and without administration of a single or a combination of individual prostaglandin receptor antagonists. The results from this study will give a clear indication of the distinctive actions of prostamides versus prostaglandins.

Given an outcome where prostamides continue to achieve activity in the presence of prostaglandin antagonists, the effectiveness of our covalent probes can be studied. Methods similar to those described in this chapter can be used to test the binding of the covalent probes and to isolate the proteins to which they bind. Employing bifunctional probes, protein

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purification techniques, and mass spectrometry, the proteins can be separated and

identified. Furthermore, incubation of the tissue with the prostaglandin receptor antagonists followed by treatment with the covalent probes may aid in the selectivity of covalent bond formation mainly with proteins distinct from the prostaglandin receptors.

In concomitance with the functional tissue studies, fluorescence confocal microscopy could be a powerful tool for identifying the diverse actions of the prostamides. Using a method similar to those by Woodword’s group discussed in Section 1.4.1.2, cells in the tissue with which the prostamide E2 and prostaglandin E2 act as signaling lipids, release

Ca2+ and can be identified by fluorescence.31 If exclusively diverse cells are activated by prostamide and prostaglandin, a strong case can be made for distinction between the prostamide and prostaglandin signaling pathways.

4.4 EP Receptor Splice Variants as the Putative Prostamide Receptor

As discussed in Section 1.4.3, strong evidence exists that the lowering of intraocular pressure by prostamide F2α glaucoma drugs such as bimatoprost, can be

attributed mainly to binding at a splice variant of the FP receptor.32 This presents the

possibility that prostamide E2 acts in an analogous fashion at splice variants of the EP receptors. Perhaps it is the EP receptor splice variants with which the prostamides can achieve their diverse physiological profiles. Investigation of the literature reveals that the genes that encode prostaglandin E receptor subtypes EP1 and EP3 also encode splice

variants for these two receptors.5 To clarify the distinction between subtypes and splice variants, the two terms are defined here. Subtypes of a given receptor preferentially recognize the same ligand, in this case PGE2, but are encoded by different genes, may be distributed in various tissues and may exert varying effects. The amino acid sequences of

receptor subtypes are diverse due to differences in the sequences of the genes that encode

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them. Alternatively, splice variants of a receptor are variations of a single receptor subtype

produced by alternative splicing during encoding by the same gene. The differences in

amino acid sequence entail truncation and result from deletions during mRNA splicing.

Splice variants are distributed within the same tissues as the parent receptor; however they

may still exert diverse effects and, importantly, may be selective for certain ligands.

The human EP1 gene encodes two distinct splice variants, one of which has a

truncated intracellular tail. Studies have determined that this alternative spliced receptor does not couple to any signal transduction cascade when treated with prostaglandin E2.

However, this does not exclude the possibility that prostamide may incite some significant

signaling on this altEP1 receptor. The human EP3 gene encodes nine alternative splice

variants giving rise to eight distinct EP3 receptor isoforms. A study similar in design to the one performed by Woodward’s group described in Section 1.4.3, employing the EP1 and

EP3 alternatively spliced receptors and subjecting them to treatment with prostamide E2 and prostaglandin E2 may lead to an important discovery in the diverse function of prostamide.

Co-expression of the alternative splice variant proteins with the wild type receptor or with other EP receptor subtypes introduces the possibility of receptor heterodimers, constituted by altEP and EP-subtype proteins. Heterodimers incorporating a splice variant protein may negate wild type activity or may exhibit altered activity. In the former case, the altEP receptor protein co-expressed with the wild type EP receptor inhibits the signaling at the wild type receptor and thus acts as an endogenous dominant negative receptor. This was found to be the case for prostaglandin E2 signaling at altEP1-EP2 and altEP1-EP4 receptors.5 Again, this does not necessarily rule out the prospect of prostamide exhibiting activity for these same heterodimers. If prostamide can indeed act at one of the altEP-wtEP

heterodimers, this would represent an altered state of activity for the prostaglandin

receptors.

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Relatedly, an EP receptor knockout study would be extremely useful to determine the

involvement of the EP prostaglandin receptors in the physiological function of prostamide E2.

Aihara’s group determined that indeed, FP receptors were required for bimatoprost and

33 prostamide F2α to maintain ocular activity. To this point, prostamides have not been tested against an EP receptor knockout model. If prostamide E2 activity is abolished by the absence of EP receptors, it can be concluded that an intact EP receptor gene is necessary for prostamide activity. It would be unlikely that there is a diverse receptor through which prostamide E2 is acting. If however, some functional activity remains, evidence suggests

that a portion of prostamide’s actions are elicited through interaction at a yet unidentified

receptor diverse from the prostaglandin EP receptors. Unlike the FP receptor study, this could entail the knockout of each receptor individually and in combination. The results determined by this study should be in accordance with those of the EP subtype receptor antagonism studies proposed.

4.5 Further Screening of Prostamide Analogs

4.5.1 Screening on PPARγ and Other Nuclear Receptors

Screening of our compounds on receptors additional to those assayed in Roth’s lab,

in particular PPAR and other nuclear receptors, may lead to discovery of novel prostamide

interactions. Prostamide E2 has been shown to be an endogenous regulator of IL-2

6 production through interactions at the PPARγ receptor. However, prostamides have yet to be tested in screening assays against a majority of the known nuclear hormone receptors.

While the prostamide F2α analog bimatoprost has been tested and found inactive against

34 estrogen receptors ERα and ERβ and glucocorticoid receptor GR, none of the prostamides

have been subjected to screening on the progesterone PR receptor, the androgen AR

receptor, the peroxisome proliferator-activated PPARα, PPARδ and PPARγ receptors, or the

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liver X LXRα and LXRβ receptors. There are numerous other nuclear receptors, however the

selection presented here are receptors for which screening assays are readily available.

Of the nuclear receptors, PPAR receptors will likely show the greatest interaction with the prostamides due to the fact that the endogenous ligands for these receptors are fatty

35 acids and prostaglandins. PPAR receptors are found throughout the body, however PPARδ may be of the paramount interest due to its enhanced expression in the brain. PPAR

receptors function as transcription factors regulating gene expression and are essential in

cellular processes such as differentiation, metabolism, and tumor genesis.36 The potency of

prostamides to bind PPAR compared to that of the fatty acids and prostaglandins is of great

interest. Were prostamides proven to be more potent than the prostaglandins, their

physiological purpose may indeed entail the activation of the PPAR receptors. However,

potency alone does not dictate prostamide’s usefulness entirely. The role of PPAR is altered

by the precise shape of the ligand-binding domain, and thus ligands that force the binding

domain into different conformations may have opposing effects. While one ligand binds to

activate receptor function the other may repress the receptors, respectively up-regulating or

down-regulating gene expression. Thus, independently of their potency, the increase bulk of

the neutral ethanolamine head group incorporated into the prostamide structure, may

impart diverse overall activity from the prostaglandins upon binding PPAR.

4.5.2 Screening on Primary Cell Systems

In addition to standard receptor screening assays, some commercial screening labs also offer numerous assays on primary cell systems. According to promotional literature

Cerep S.A., a French-based company, offers one hundred and twelve individual cellular assays in seventeen different cell systems including, but not limited to, neural, ocular, cardiac, pulmonary, reproductive, gastrointestinal, dermal, and blood cellular systems. Of greatest interest to the study of prostamide would be the twenty four neural cell assays due

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to the increased concentration of anandamide within the neural system and perhaps the

seven ocular cell assays due to the effective lowering of intraocular pressure by

prostamides. Screening the prostamides and prostaglandins against neural cells such as

astrocytes, neurons, brain microvascular endothelial cells, brain vascular smooth muscle

cells, and meningeal cells may lead to better understanding of which cells are particularly

sensitive to activation by prostamides and whether these effects are diverse from those of

prostaglandins. Once a specific set of cells is identified as particularly responsive to

prostamide, focus can be shifted to the identification of the proteins within the cell with

which prostamides are interacting to elicit their effects. Again, covalent modification and bifunctional probes used to fish out these proteins may represent the subsequent step that leads to the isolation and identification of such proteins. While exceedingly valuable information can be gained from these screens of primary cell systems, the main drawback is the extreme expense required to carry out the testing.

4.6 Prostamide Activity in the Resolution of Inflammation

Further collaboration with Dr Serhan’s laboratory at Harvard Medical School to test our remaining prostamide analogs on in vivo PMN and monocyte concentration may lead to the identification of particularly potent or efficacious compounds (Section 3.2.1). Due to the large number of mice sacrificed and the extended effort required, this method would not be feasible for a large scale SAR study. Additionally, in vivo testing does not allow for the determination of covalent binding ability of the probes to the proteins with which they interact. For this, ex vivo tissue studies, like those describe generally in Section 4.1.1, may be more useful. Thus, a number of abdominal tissue samples can be harvested from a single mouse, treated with zymosan A to activate an inflammatory response, and used to test the effects of several prostamide probes on PMN and monocyte concentration. Once acceptable potency of a probe has been achieved within these assays, bifunctional probes can be

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employed to aid in extracting and identifying the proteins within the inflamed abdominal

tissue that bind to prostamide.

Of particular interest would be the administration of EP receptor antagonists in

conjunction with the prostamides and prostaglandins. Our results demonstrated a marked

decrease in PMN infiltration when treated with prostaglandin E2 and the 20-isothiocyanato

prostamide E2 probe, AM7645. Incubation of the inflamed abdominal tissue with ONO8711

or SC51089 would selectively antagonize the EP1 receptor and the involvement of this particular EP receptor in the prostamide-induced clearance of PMN can be ascertained. If

PMN levels remain unchanged when treated with EP1 antagonist, the EP1 receptor is not engaged in PMN regulation by prostamide. However, if EP1 antagonist reverses all or some

of the PMN decrease incited by prostamide, it is likely that this specific receptor is indeed

involved in PMN regulation by prostamide. Likewise, AH6809 and PF04418948 antagonists

can be administered to assess the involvement of EP2, DG041 and CM9 antagonists for EP3,

and AH23848B and CJ023423 antagonists for EP4. This study would essentially confirm or deny the involvement the prostaglandin receptors in the facilitation of inflammatory resolution by prostamide E2 and prostaglandin E2. If the prostaglandin antagonists do not reverse prostamide’s clearance of PMN the results would represent a substantial discovery in the understanding of prostamide function. There is indeed a distinct target diverse from the prostaglandins with which prostamide is interacting and eliciting pro-resolving effects.

4.7 Suppression of the Immune System by Prostamide versus Anandamide

Additional exploration into the immunosuppressive abilities of prostamide may lead to intriguing discoveries of its involvement in immune system regulation. For example,

anandamide has long been associated with immunosuppression due to its activation of the

CB2 receptor; however it is unknown whether the metabolic conversion of anandamide to

prostamide represents a termination of immunosuppressive effects. Because the first set of

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results presented in Section 3.3 demonstrates that prostamide E2 and its analogs do

possess the ability to suppress the immune system response, metabolism of anandamide to

prostamide may not entail the complete loss of immunosuppressive effect. Perhaps some of

anandamide’s immunosuppression can be attributed to its prostamide metabolite at a target

disparate from the CB2 receptor. To test this hypothesis, the PFC and MLR studies which monitor lymphocyte levels produced in response to antigen can be repeated with anandamide in the presence and absence of a COX inhibitor. If suppression of the immune response by anandamide in the presence of COX inhibitor is identical to that without COX inhibitor, it is likely that anandamide’s immunosuppressive activity can be attributed entirely to activity at the CB2 receptor. If however, COX inhibition alters the potency or efficacy of

the established immune response by anandamide, prostamide may indeed play a role in the anandamide immunosuppression. Comparison of anandamide potency to that of prostamide

E2 may give an indication of the physiological function of conversion from anandamide to

prostamide.

Repetition of the immunology experiments at higher concentrations may give more

accurate results as THC’s optimal range for immunosuppression is 10-32 µM and the

prostamide analogs were tested up to 10 µM. Prostaglandin E2 should also be tested to compare the results with those of prostamide E2. Again, this may lead to a distinction between prostamide and prostaglandin action. Additionally, attempting these assays in the presence of the individual EP receptor agonists will give an indication if any of the prostaglandin receptors are responsible for prostamide’s suppression of the immune response.

4.8 Synthesis of Additional Prostamide Analogs for Biological Testing

Development of more potent prostamide probes is contingent upon establishing an effective assay that is selectively responsive to prostamide. The collaborative studies

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proposed above may lead to such an assay allowing for the design of a large library of compounds to test for unique prostamide response and eventually to aid in isolation and identification of target proteins. The design of future analogs should focus mainly on functionalization of the ω-tail to increase potency and/or enhance covalent bonding and of the α-head to increase metabolic stability and incorporate labels essential for isolation and identification of the covalently modified proteins.

4.8.1 Design and Proposed Synthesis of ω-Tail Modifications to Enhance Binding Affinity

A nearly limitless number of modifications can be made on either end of the central prostamide backbone. Figure 2.16, shown again here, illustrates a sampling of the alterations to the tail moiety that are easily available synthetically. Ethers, carbamates, esters, alkenes, amides, imides and epoxides are all achievable in relatively few steps from the free alcohol. Select analogs can be designed based on optimization of an SAR study depending on the system. An initial SAR focuses on crude changes in size and polarity to gain an overall understanding of which types of modifications are tolerable for binding, and once this knowledge is gained, further fine tuning of the analogs to the binding site can commence. Thus, the first set of analogs should incorporate a large, medium, and small functional group such as adamantyl, t-butyl, and methyl groups, respectively, in order to ascertain the steric restrictions of the binding site. For example, if a large group is easily accommodated, there is most likely a large binding pocket in which this group can easily fit.

Additionally, hydrogen bond donor/acceptors such as hydroxyls, thiols, amines, or carboxyls should be incorporated to probe for important hydrogen bond or salt bridge interactions within the binding pocket. Charged amino acid residues capable of forming salt bridges and hydrogen bonds include arginine, histidine, aspartic acid, and glutamic acid. Neutral serine and asparagine can also serve as hydrogen bond donors and acceptors. Electrostatic

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Figure 2.16 Synthetic directions allowed by the key intermediate alcohol 20. (LG = Leaving group, Ox = Oxidant)

interactions such as these have an enormous effect on binding potency. Finally, overlapping

somewhat with the previous two areas, is the introduction of hydrophilic or hydrophobic groups to optimize like interfaces in the binding site. If there exists a hydrophobic pocket made up of hydrophobic residues such as phenylalanine, isoleucine, or valine within the binding site, ligand moieties containing hydrophobic groups such as phenyl or alkyl groups will bind more readily. A hydrophilic area within the binding site may constitute an opening or channel exposed to cellular fluid or a pocket composed of polar amino acid residues such as lysine, arginine, or aspartic acid. In this case, analogs containing polar moieties like hydroxyls, thiols, and carboxyls should cause the compounds to bind more potently.

The synthesis of further tail modified prostamide analogs would be similar to that presented for the covalent probes in Scheme 2.2. From the free hydroxyl 20, several

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derivatives can be arrived at through a few additional steps. (Figure 2.16) For example, an

array of carbamates can be produced upon reaction of the primary alcohol with almost any

available isocyanate. Conversion of the hydroxyl to a leaving group such as a tosylate would

allow for SN2 substitution reactions with nucleophiles such as the cyano group, primary amines, thiols, halides or other nucleophiles. Addition of a leaving group can also aid in elimination of the alcohol to a terminal alkene. The alkene can then be taken for further reaction including addition across the double bond or Wittig-type reactions. Oxidation of the alcohol to the aldehyde opens up a number of possibilities including Aldol condensations,

amine condensations and Wittig reactions with various carbonyl, amine, and alkene reagents, respectively. Further oxidation to give the carboxylic acid introduces the possibility for numerous ester and amide coupling reactions. Finally, a Williamson ether reaction can be used to incorporate various ether substituents at the tail position. All of these synthetic directions are easily attainable from our intermediate, and expansion of a library of prostamide analogs integrating the alterations in size and electron density discussed previously can be carried out for a thorough SAR study.

For example, to investigate the size of the active site binding pocket without substantially altering any electrostatic interactions, the methyl, t-butyl, and adamantyl groups can be introduced as ethers. (Figure 4.7) Due to the overall hydrophobicity of all of these groups, an SAR incorporating them may lead to an overall model of the hydrophobic/hydrophilic requirements for binding. Introduction of phenyl ethers using similar methods will give more information pertaining to the hydrophobicity of the binding site including optimization of Van der Wahl’s interactions. Hydrophilicity can be further gauged using various polar group substitutions. If thiol and amino tail groups synthesized in

Figure 4.8 bind well, the indication is that a hydrophilic pocket is present or that there is an important hydrogen bond interaction. Oxidation of the alcohol to a carboxylic acid and

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Figure 4.7 Use of Williamson ether reactions to introduce various sizes of hydrophobic substituents for SAR study of prostamide binding.

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Figure 4.8 Proposed synthesis of carboxylic acid, thiol, and amino tail modified prostamide analogs for SAR study of prostamide binding.

Once a better understanding of the structural modifications required for potent binding has been established through the SAR results, the covalent isothiocyanato and azido modifications can be reintroduced. With highly potent covalent probes in hand, the process of isolating proteins with which the prostamide analogs bind can begin. For that, bifunctional probes incorporating radiolabels and biotin labels can be designed for isolation and identification of the proteins.

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4.8.2 α-Head Modifications to Increase Metabolic Stability and Incorporate Radio- and Biotin Labels

Modifications of the head group should incorporate the amide constituent necessary

to represent prostamide structure along with substitutions to increase metabolic stability

and the ability to incorporate labeling functionality. The covalent probes described within

this dissertation suitably integrate methyl and cyclopentyl groups previously shown to

increase anandamide stability for this purpose. Design of future analogs should attempt to

continue including substitutions to enhance stability without adverse effects on potency.

Thus R-methyl substitution at the 1-position of the ethanolamide should be included to

protect from metabolism by esterase and amidase enzymes.

4.8.2.1 Design and Proposed Synthesis of Radiolabeled Prostamide Analogs

Once potent, covalent probes are obtained from detailed SAR study, the next step is

designing bifunctional probes integrating both a covalent-bond forming modification and a

label with which to isolate and identify the ligand-bound protein. This can be done using a

radiolabel such as 125I or through a biotin label. As discussed in Section 4.1.2, radiolabeling allows for facilitated identification of the ligand-bound proteins purified by PAGE gel separation and for use as radiotracers in PET and SPECT imaging. The amide head group is

ideal for the inclusion of a radiolabel as placing the radiolabel here ensures that the bound

ligand is the entire prostamide and not the product of hydrolysis to prostaglandin. Key to

the synthesis of radiolabeled probes is the ability to attach the radioisotope as the last step

prior to biological testing. This reduces exposure to radioactivity and ensures that

substantial radioactivity remains for the biological testing or radiolabeled imaging.

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Figure 4.9 Proposed example for the synthesis of radioiodinated prostamide probes by coupling to bromine-carrying amines in the amidation step and by subsequently replacing the bromine with iodine in a Finkelstein reaction.

Incorporation of a radioactive iodine in the ultimate step can be achieved using an overall synthesis similar to that of the current covalent probes but incorporating a bromine group within the coupled amide substituent. (Figure 4.9) Conversion of the bromine to the radioactive iodine using a Finkelstein reaction allows exchange labeling with radiolabeled sodium iodide to be done in the final step. Thus, as long as the amine to which the carboxylic acid 23 is coupled contains a bromine or a moiety that can easily be converted to a bromine, radioiodination can easily be achieved. For example, a hydroxyl can be modified into a bromide by phosphorus tribromide and alkenes can undergo addition of bromine by

N-Bromosuccinimide (NBS). (Figure 4.10) Additionally, there are a plethora of commercially available primary amine compounds containing one or more bromine substitutions that could be useful for easy expansion of the library. (Figure 4.11)

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Figure 4.10 Synthesis of brominated primary amines for coupling and subsequent exchange labeling with radioactive iodide.

Figure 4.11 Examples of possible radioiodinated analogs of the covalent prostamide probes to form bifunctional radiolabeled probes.

4.8.2.2 Design and Proposed Synthesis of Biotinylated Prostamide Analogs

Alternatively, introduction of a biotin label in place of the radiolabel would allow for

direct extraction of proteins to which the prostamide probes bind through the use of an

avidin-coated solid bead. (Section 4.1.2.3) The key to a successful biotinylated probe is the

inclusion of a sufficiently long linker between the ligand itself and the biotin moiety. The

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isolation process essentially attaches the binding site of prostamide-bound protein to the binding site of the avidin protein. Due to the large relative size of proteins, the bifunctional probes require ample distance between the portions of the ligands that bind to each protein.

Even more so than the radioligand studies, this method requires successful covalent binding of the probes to the receptor proteins. Thus the length of the chain connecting the biotin functionality to the prostamide probe may play an important role in the success of the affinity purification process. There are a number of biotin labeling reagents available containing linkers of various lengths and tail functionalities to couple to the ligand. Often the linker is made up of polyethylene glycol, and is frequently comprised of four mer units.

Figure 4.12 The Click Reaction forms a triazole from an azide and an alkyne and utilizes copper (I) as a catalyst.

A suitably straightforward method to attach a biotin label is through the use of the

Click reaction. The Click involves the formation of a triazole from an azide and an alkyne.

(Figure 4.12) The reaction is catalyzed by copper (I), and it is frequently used in biochemistry to bring together two distinct ligands or proteins into one bifunctional complex.

The azide and alkyne functionalities can be used interchangeably, thus it is relatively arbitrary which of the two should be incorporated into the ligand. Commercially available

biotin labeling reagents are often available both with an azide and with an alkyne. An

example of biotin labeling one prostamide probe with a commercial reagent using the Click

reaction is illustrated in Figure 4.13.

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Figure 4.13 Proposed example for the synthesis of biotinylated prostamide probes by Click reaction from azide and alkyne functionalities.

The Click reaction would not be performed on the azido probes due to the obvious selectivity issues. Instead, a Sonogashira coupling with a halogenated biotin-linker would lead to successful synthesis of the bifunctional probe. Synthesis of a halogenated linker from an ethylene glycol starting material followed by coupling to biotin is proposed in

Figure 4.14. With the iodinated biotin complex in hand, it can be coupled to the prostamide probe using Sonogashira conditions.

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Figure 4.14 Proposed synthesis of biotinylated 20-azido prostamide probes

The alkyne head group alone could lead to a number of possible biochemistry

studies. For example, following incubation of the prostamide probe with the target receptor

proteins, a fluorophore can be introduced through a Click reaction. The result would be fluorescently labeled proteins that could be observed under a microscope and quantified under a fluorometer. In fact, it may prove to be more effective for the affinity purification to introduce only the alkyne head group to the ligand and incubate the ligand with the target proteins. Subsequently, an avidin-coated bead bound with the biotin-linker-azide reagent can be introduced and the Click reaction initiated to form the anchored prostamide-binding protein. This method may be beneficial due to the hypothetical increase in the initial binding of the prostamide probe to the protein of interest during incubation. In addition, we could test our azido covalent probes with this method due to the inaccessibility of the azido group that is theoretically covalently bound to the receptor protein. However, the Click reaction itself may prove to be more challenging using this method due to the steric bulk of the two

-203- proteins coming together to form the linkage. A third option would be to complete the experiment in three steps. The initial step would be to incubate the sample of interest with the prostamide probe as before but the following step would involve the Click reaction of the biotin-linker-azide alone. Finally, introduction of the avidin-coated bead to bind the biotin would successfully anchor the target proteins to the bead for further purification.

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12. Kawada, K.; Dolence, E. K.; Morita, H.; Kometani, T.; Watt, D. S.; Balapure, A.; Fitz, T. A.; Orlicky, D. J.; Gerschenson, L. E., Prostaglandin photoaffinity probes: synthesis and biological activity of azide-substituted 16-phenoxy- and 17-phenyl-PGF2.alpha. prostaglandins. J. Med. Chem. 1989, 32, 256.

13. Watanabe, K.; Kawamori, T.; Nakatsugi, S.; Ohta, T.; Ohuchida, S.; Yamamoto, H.; Maruyama, T.; Kondo, K.; Ushikubi, F.; Narumiya, S.; Sugimura, T.; Wakabayashi, K., Role

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of the prostaglandin E receptor subtype EP1 in colon carcinogenesis. Cancer Res. 1999, 59, 5093.

14. Hallinan, E. A.; Hagen, T. J.; Husa, R. K.; Tsymbalov, S.; Rao, S. N.; vanHoeck, J. P.; Rafferty, M. F.; Stapelfeld, A.; Savage, M. A.; Reichman, M., N-Substituted dibenzoxazepines as analgesic PGE2 antagonists. J. Med. Chem. 1993, 36, 3293.

15. Kiriyama, M.; Ushikubi, F.; Kobayashi, T.; Hirata, M.; Sugimoto, Y.; Narumiya, S., Ligand binding specificities of the eight types and subtypes of the mouse prostanoid receptors expressed in Chinese hamster ovary cells. Br. J. Pharmacol. 1997, 122, 217.

16. af Forselles, K. J.; Root, J.; Clarke, T.; Davey, D.; Aughton, K.; Dack, K.; Pullen, N., In vitro and in vivo characterization of PF-04418948, a novel, potent and selective prostaglandin EP2 receptor antagonist. Br. J. Pharmacol. 2011, 164, 1847.

17. Heptinstall, S.; Espinosa, D. I.; Manolopoulos, P.; Glenn, J. R.; White, A. E.; Johnson, A.; Dovlatova, N.; Fox, S. C.; May, J. A.; Hermann, D.; Magnusson, O.; Stefansson, K.; Hartman, D.; Gurney, M., DG-041 inhibits the EP3 prostanoid receptor. - A new target for inhibition of platelet function in atherothrombotic disease. Platelets 2008, 19, 605.

18. Jugus, M. J.; Jaworski, J. P.; Patra, P. B.; Jin, J.; Morrow, D. M.; Laping, N. J.; Edwards, R. M.; Thorneloe, K. S., Dual modulation of urinary bladder activity and urine flow by prostanoid EP3 receptors in the conscious rat. Br. J. Pharmacol. 2009, 158, 372.

19. Coleman, R. A.; Grix, S. P.; Head, S. A.; Louttit, J. B.; Mallett, A.; Sheldrick, R. L. G., A novel inhibitory prostanoid receptor in piglet saphenous vein. Prostaglandins 1994, 47, 151.

20. Murase, A.; Nakao, K.; Takada, J., Characterization of Binding Affinity of CJ-023,423 for Human Prostanoid EP4 Receptor. Pharmacology 2008, 82, 10.

21. Takahashi, H.; Higuchi, M.; Suhara, T., The Role of Extrastriatal Dopamine D2 Receptors in Schizophrenia. Biol. Psychiatry 2006, 59, 919.

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24. Wager, K. M.; Jones, G. B., Radio-iodination methods for the production of SPECT imaging agents. Curr. Radiopharm. 2010, 3, 37.

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26. Corbett, J. R., Fatty acids for myocardial imaging. Semin Nucl Med 1999, 29, 237.

27. Knapp, F. F., Jr.; Kropp, J.; Franken, P. R.; Visser, F. C.; Sloof, G. W.; Eisenhut, M.; Yamamichi, Y.; Shirakami, Y.; Kusuoka, H.; Nishimura, T., Pharmacokinetics of radioiodinated fatty acid myocardial imaging agents in animal models and human studies. Q J Nucl Med 1996, 40, 252.

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28. Sinz, A., Isotope-labeled photoaffinity reagents and mass spectrometry to identify protein-ligand interactions. Angew. Chem., Int. Ed. 2007, 46, 660.

29. Roth, A. F.; Wan, J.; Green, W. N.; Yates, J. R.; Davis, N. G., Proteomic identification of palmitoylated proteins. Methods (San Diego, CA, U. S.) 2006, 40, 135.

30. Kaschani, F.; Gu, C.; Niessen, S.; Hoover, H.; Cravatt, B. F.; van, d. H. R. A. L., Diversity of serine hydrolase activities of unchallenged and Botrytis-infected Arabidopsis thaliana. Mol. Cell. Proteomics 2009, 8, 1082.

31. Spada, C.; Krauss, A.; Woodward, D.; Chen, J.; Protzman, C.; Nieves, A.; Wheeler, L.; Scott, D.; Sachs, G., Bimatoprost and prostaglandin F selectively stimulate intracellular calcium signaling in different cat iris sphincter cells. Exp. Eye Res. 2005, 80, 135.

32. Liang, Y.; Woodward, D. F.; Guzman, V. M.; Li, C.; Scott, D. F.; Wang, J. W.; Wheeler, L. A.; Garst, M. E.; Landsverk, K.; Sachs, G.; Krauss, A. H. P.; Cornell, C.; Martos, J.; Pettit, S.; Fliri, H., Identification and pharmacological characterization of the prostaglandin FP receptor and FP receptor variant complexes. Br. J. Pharmacol. 2008, 154, 1079.

33. Ota, T.; Aihara, M.; Narumiya, S.; Araie, M., The effects of prostaglandin analogues on IOP in prostanoid FP-receptor-deficient mice. Invest Ophthalmol Vis Sci 2005, 46, 4159.

34. Woodward, D. F.; Liang, Y.; Krauss, A. H. P., Prostamides (prostaglandin- ethanolamides) and their pharmacology. Br. J. Pharmacol. 2008, 153, 410.

35. Harmon, G. S.; Lam, M. T.; Glass, C. K., PPARs and Lipid Ligands in Inflammation and Metabolism. Chem. Rev. (Washington, DC, U. S.) 2011, 111, 6321.

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Chapter 5: Inhibition of Fatty Acid Amide Hydrolase by a Novel Class of Prostamide Intermediates

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5.1 FAAH Enzyme Structure and Function

Fatty Acid Amide Hydrolase (FAAH) is the enzyme responsible for the hydrolysis of

amide bonds in fatty acid amides (FAAs) including the N-acyl ethanolamines, N-arachidonoyl ethanolamide (anandamide, AEA),1 N-stearoyl ethanolamide (SEA), N-palmitoyl

ethanolamide (PEA),2 and N-oleoyl ethanolamide (OEA),3 and the fatty acid primary amide, oleamide.4,5 (Figure 5.1) FAAs are associated with a wide variety of biological actions. For

example, AEA is associated mainly with analgesia,6 PEA with anti-inflammation,7 OEA with

satiation signaling,8 and oleamide with sleep induction.4 The various actions of these

bioactive lipids are terminated when their amide bonds are cleaved by the action of the site

specific FAAH.9-11 Most notably, anandamide’s cannabinergic signaling is degraded by FAAH catabolism to arachidonic acid. This simple conversion of amide to carboxylic acid is sufficient to produce a loss of activity at CB1 and constitutes the main metabolic breakdown of anandamide.

Figure 5.1 Known FAAH substrates

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A

B C

Figure 5.2 (A) Representation of the quaternary crystal structure of rFAAH shown bound to the membrane, (B) View of MAP within the binding pocket of rFAAH highlighting the membrane binding alpha helices, α18 and α19, and the cytosolic port for expulsion of amine and introduction of water, (C) Orientation of MAP bound to the active site of rFAAH illustrating the Ser-Ser-Lys catalytic triad and the nearby residues that contribute to binding.12

FAAH, an integral membrane protein, is the only mammalian member of the amidase signature (AS) class of enzymes, and is anchored to the interior of the cell by a single N- terminal transmembrane domain.9,13 FAAH’s crystal structure comprises a dimeric enzyme

and includes two hydrophobic alpha helices, α18 and α19, that integrate into the membrane.4,14 (Figure 5.2) The α18 and α19 alpha helices act as hydrophobic caps at the

entrance channel that leads to the enzyme’s active site, and the adjacent positioning of the

active site to the membrane is ideal for the admission of lipid soluble substrates such as the

fatty acid amides.12 While this entrance area is mostly lined with hydrophobic side-chains, nearby charged amino acids, Asp403 and Arg486, may play a role in orientation of the substrate towards the active site by interacting with the polar amide head group. This

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overall arrangement is similar to that of the COX enzymes which act on similarly

hydrophobic arachidonic acid.15 A second channel within the FAAH tertiary structure leads

out to the cytosol from the active site and may represent the exit route through which the

polar amine groups, cleaved from the fatty acid amide, can be released, and through which water, required for the hydrolysis may be provided.

Amide Hydrolysis

Ester Hydrolysis from the enzyme

Figure 5.3 Representation of FAAH’s catalytic mechanism

Once in the active site, the fatty acid amide is acted on by a nucleophilic serine-

241.16 The nucleophilicity of Ser-241 arises from an unusual Ser-Ser-Lys catalytic triad

which is characteristic of the AS class of enzymes.12,16-19 (Figure 5.3) The Lys142 accepts a

hydrogen bond from Ser217, and the Ser217, in turn, accepts a hydrogen bond from

Ser241, thus producing a nucleophilic oxygen on Ser241 capable of attacking the carbonyl

of the fatty acid amide. The amide nitrogen then forms a bond with the hydrogen from

Ser241 and, when the carbonyl is reestablished, leaves as an amine. With the fatty acid

now esterified to the FAAH itself, it must be cleaved from the active site. A molecule of

water enters the site through the cytosolic channel and undergoes a similar hydrogen

bonding cascade with Ser217 and Lys142 to release the fatty acid product.

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5.2 Inhibition of FAAH

As previously stated, FAAH’s most notable biological action is the degradation of the

endocannabinoid signaling pathway achieved by the chemical deactivation of anandamide.

Thus it follows that inhibition of the FAAH enzyme would increase endocannabinoid tone and

give similar physiological effects to those associated with activation of CB1. Indeed, FAAH

knockout mice have shown elevated levels of anandamide, PEA, OEA, and oleamide as well

as the overall analgesic, anxiolytic, antidepressant, sleep-enhancing and anti-inflammatory

phenotypes related to agonism by these compounds.20-24 Interestingly, the undesirable side- effects of direct CB1 agonism on motility, weight gain, and body temperature are not evident in FAAH knockout mice.22,25 As a result of FAAH’s ability to indirectly increase

activation of the cannabinoid receptors without these side-effects, it has become an

attractive target for treatment of neuropathic pain.

5.2.1 Substrate-Based, Nonselective Inhibitors

Figure 5.4 Structures and potencies of the first identified class of nonselective FAAH inhibitors

FAAH inhibitors can be grouped into two distinct classifications: activated carbonyl

compounds that form reversible hemiketals with the active site Ser241, and carbamate or urea compounds that form irreversible covalent bonds with the active site Ser241. The earliest FAAH inhibitors were derived from the first endogenous compound to exhibit FAAH inhibition, 2-octyl α-bromoacetoacetate, and modeled after the oleamide substrates.26-28

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(Figure 5.4) These compounds maintain all that is required for FAAH inhibition – an

electron-poor carbonyl carbon and a long hydrophobic chain. Following extensive SAR study, it is evident that activated carbonyl compounds having stronger electron-withdrawing groups, such as the trifluoromethylketone, give more potent inhibition.28-32 Additionally, the

phenhexyl group is found to be a well-tolerated replacement for the long hydrophobic chain

by imparting the lipophilicity required for binding the active site.29 Although the compounds

within this initial class are very potent for FAAH inhibition, they exhibit very low selectivity

for FAAH. They inhibit phospholipase A (PLA),33 platelet-activating factor acetylhydrolase

(PAFAH),34 and monoacylglycerol lipase (MGL).35 In fact, they give inhibition of triacylglycerol hydrolase (TGH) and serine hydrolase KIAA1363 at even greater potency and are considered reversible inhibitors of the entire serine hydrolase class of enzymes.30

5.2.2 Selective, Reversible Inhibitors

Figure 5.5 Structures and potencies of the first, selective FAAH inhibitors

The first, selective FAAH inhibitors to be developed were the potent and reversible α- ketoheterocycle analogs.36,37 (Figure 5.5) SAR study led to the determination that oxazole

is the most potent monocyclic substituent tested, while the bicyclic oxazolopyridines give

the overall most exceptionally potent FAAH inhibition.36,37 Incorporation of the phenyl tail in

place of the long oleoyl chain produced a FAAH inhibitor with excellent potency and

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enhanced selectivity for FAAH over TGH and KIAA1363 (>100-fold). Further SAR

development yielded OL-135, the most well studied reversible, competitive, and selective

FAAH inhibitor to date.38 Numerous analogs of OL-135 have since been designed to increase selectivity, potency, and solubility. Inhibition by reversible FAAH inhibitors requires an electron-poor carbonyl carbon to act as an electrophile for nucleophilic attack by the activated Ser241 in the Ser-Ser-Lys catalytic triad to form a hemiketal.39,40 (Figure 5.6)

Figure 5.6 Mechanism for the reversible inhibition of FAAH

A plethora of both competitive and noncompetitive reversible FAAH inhibitors have

been designed as therapeutics by a number of pharmaceutical companies. Several different

classes of reversible FAAH inhibitors have arisen including substituted thiohydantoins and

imidazolidinediones,41,42 oxime carbamates,43 enol carbamates,44 benzothiazole,45 cylic

ureas,46,47 benzoxazole,48,49 aminopyrimidine,50 oxadiazolylphenylboronic acids,51,52 α- ketoamide,53,54 and β-lactams.55 (Figure 5.7) Many of these compounds appear to bear the

structural elements required for irreversible inhibition, and in some cases it is difficult to

imagine that these inhibitors do not acylate the active site Ser241. However, the results of

their biological assays demonstrate that they are in fact reversible as there is no increase in

their potency over an extended incubation time and they are dializable from the enzyme.

Several of the reversible FAAH inhibitors have undergone and are currently undergoing

Phase 1/2 clinical study for the treatment of pain, inflammation, and sleep disorders,

however none are presently being marketed as drugs.51,52,56

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Figure 5.7 Several of the current industry-leading reversible FAAH inhibitors as representatives of various classes of FAAH inhibitors.

5.2.3 Nonselective, Irreversible Inhibitors

Figure 5.8 Structures methylarachidonylfluorophosphonate (MAFP) Phenylmethylsulfonylfluoride (PMSF) – two commonly used irreversible inhibitors

Phenylmethylsulfonylfluoride (PMSF) and methylarachidonylfluorophosphonate

(MAFP) are two irreversible, nonselective serine protease inhibitors commonly used in studies where FAAH inhibition is necessary.12,57 (Figure 5.8) PMSF is used broadly in

biochemical protein preparations to inhibit serine proteases from digestion of proteins

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following cell lysis, and in bioassays to inhibit the hydrolysis of designed fatty acid amide analogs prior to their activity at receptors like CB1. MAFP, on the other hand, is more commonly used in X-ray crystallography studies of FAAH, as it possesses the acyl arachidonyl chain of the substrate, anandamide, and the electrophilic, fluorinated carbonyl optimal for covalent binding to Ser241. Thus, MAFP is ideal for the study of the active site orientation of anandamide when bound to substrate. The activated carbonyl carbon of these two irreversible inhibitors is prone to nucleophilic attack by the active site serine and maintains this bond by expulsion of the fluorine leaving group.

Figure 5.9 Structures of the initial lead compound, carbaryl, and the current irreversible FAAH inhibitors, URB524 and URB597

The earliest irreversible FAAH inhibitors disclosed were derived from modifications of

the acetylcholinesterase inhibitor (AChEI), carbaryl.58 (Figure 5.9) The first in an ever- expanding class of o-aryl carbamate irreversible inhibitors, URB524 and URB597 are potent, noncompetitive, irreversible FAAH inhibitors that exhibit poor selectivity for FAAH.25,59

Studies determined the o-aryl carbamates to be irreversible based on their increased potency with extended incubation times and due to the fact that they are nondializable from the enzyme.25,60 Crystallographic results illustrate that the carbamate portion of the molecule binds covalently to the Ser241, while the phenolic moiety serves as the leaving group.61 (Figure 5.10) Second generation o-aryl carbamates have been developed by

several pharmaceutical companies and offer greater metabolic stability and enhanced

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Figure 5.10 Mechanism for the irreversible inhibition of FAAH by (A) carbamates and (B) ureas.

Figure 5.11 Current industry-leading carbamate-based irreversible FAAH inhibitors.

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Figure 5.12 Current industry-leading urea-based irreversible FAAH inhibitors.

Aryl ureas are another class of irreversible FAAH inhibitors being pursued by several

groups. (Figure 5.12) One of the earliest is Lilly’s tetrazole-based LY-2183240.71,72

Triazolopyridine carboxamides and ureas,73,74 piperazine aryl ureas,75 benzothiophene piperazine ureas,76 and chiral azetidines77 have also been developed as effective FAAH/MGL

inhibitors. To date, the most broadly studied and successful irreversible FAAH inhibitors in

regards to potency and selectivity for hFAAH, are Pfizer’s prototype PF-622 and PF-750.78

Extensive SAR on these compounds led to the exceptionally potent and selective PF-

04457845.79

5.3 Rat FAAH vs Human FAAH: Differences in Inhibitor Sensitivity Profiles

Rat,9 human,10 mouse,10 and pig80 orthologues of FAAH have been cloned and share over 80% sequence identity. rFAAH is the most common homologue employed in FAAH studies due to its relatively high yielding expression and enhanced stability when compared to the other species of FAAH. In fact, because rFAAH has been the standard enzyme of

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choice, reports of “FAAH inhibition” prior to 2011 give only the inhibition activity of the

compounds tested on rFAAH. While the homologous FAAH enzymes share a similar general

selectivity, rFAAH and hFAAH have more recently been shown to exhibit marked differences

in their inhibitor sensitivity.81 For example, the well-known, irreversible inhibitors PF-750

and PF-622 are 7.6-fold and 4.0-fold more potent, respectively, in hFAAH than rFAAH.

(Table 5.1) Conversely, the familiar, reversible inhibitor OL-135 exhibits a 4.4-fold greater

potency in rFAAH than hFAAH. Thus, some portion of the 18% disparity between rat and

human FAAH sequence is concentrated within the active site and is essential for selectivity.

Table 5.1 Comparison of the affinities of well-known inhibitors for rat FAAH, human FAAH, and humanized rat FAAH. Inhibitor Structure hFAAH rFAAH h/r Ratio FAAH h/rFAAH to rFAAH

PF-750 O Kinact/Ki Kinact/Ki Kinact/Ki Potency N = 791 = 104 = 528 ratio = 7.6 N N H ± 34 ± 14 ± 53 M-1s-1 M-1s-1 M-1s-1

PF-622 O Kinact/Ki Kinact/Ki Kinact/Ki Potency = 621 = 154 = 623 ratio = 4.0 N N H ± 130 ± 67 ± 250 N -1 -1 -1 -1 -1 -1 N M s M s M s

OL-135 O IC50 = IC50 = IC50 = IC50 ratio = O N 208 47.3 420 4.4* ± 35 ± 2.9 ± 15 N nM nM nM

*Expressed as a ratio of IC50 values (reversible inhibition), not a ratio of Kinact/Ki (irreversible inhibition). The value 4.4 indicates that OL-135 is 4.4-fold more potent in rFAAH than in hFAAH.81

Indeed, one study has reported a disparity in several residues of the rFAAH and hFAAH active sites possibly leading to this inhibitor selectivity.81 To date, only the crystal

structure of rat FAAH has been elucidated (Figure 5.2),12 as human FAAH has proven a challenge to express in high yield in recombinant systems and is relatively unstable and prone to aggregation. A group at Pfizer attempted an approximation of the hFAAH active site crystal structure using site-directed mutagenesis of rFAAH to create a humanized rat

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FAAH protein.81 As such, the expression levels and stability could be optimized close to that

of rFAAH, while the active site would maintain a residue sequence identical to that of

hFAAH. Active-site residues identified from the rFAAH crystal structure were compared to

the sequence of hFAAH residues, and six dissimilar amino acids were identified. All of these

residues were found to be in the proximity of the acyl arachidonyl chain of the rFAAH crystal

structure bound to the irreversible covalent inhibitor methylarachidonylphosphonate (MAP).

Upon mutation of rFAAH’s L192, F194, A377, S435, I491, and V495 to hFAAH’s F192, Y194,

T377, N435, V491, and M495, a humanized rat FAAH (h/rFAAH) protein was established.

The expression levels were improved to approximately 10mg of protein per liter of culture

versus hFAAH’s approximately 1mg of protein per liter of culture. Furthermore, rFAAH,

hFAAH, and h/rFAAH exhibit similar catalytic efficiencies. The inhibitor sensitivity profile for

h/rFAAH is similar to that of hFAAH (Table 5.1), verifying that it is a good approximation of the hFAAH enzyme.

Most importantly, enhanced stability allowed for determination of the h/rFAAH crystal structure bound to the covalent inhibitor PF-750.81 The overall fold of h/rFAAH was found to

be nearly identical to that of rFAAH. However, differences in the active site interactions with

PF-750 may explain the disparity in inhibitor sensitivity between rat and human FAAHs.

Figure 5.13 gives the comparative crystal structures of PF-750 (purple) bound to the active

site of h/rFAAH (blue residues) and to the active site of rFAAH (yellow residues). All six of

the amino acid adaptations within the active site are represented. The F381 and F192 of

h/rFAAH exhibit favorable CH-π interactions with the quinoline ring of PF-750. Because

rFAAH lacks the F192 and contains a L192 in its place, there is a reduction in affinity of

rFAAH for PF-750. Additionally, the greater steric demand of rFAAH’s I491 compared to

h/rFAAH’s V491 means that PF-750 may be too large to be easily accommodated into the

active site and this also could contribute to the higher potency of PF-750 for hFAAH over

rFAAH.

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Figure 5.13 Comparative crystal structure of PF-750 (purple structure) bound to the active site of rFAAH (yellow residues) and to the active site of h/rFAAH FAAH (blue residues).81

Although the h/rFAAH enzyme has given valuable insight into the possible

interactions of hFAAH with inhibitors and the disparity in inhibitor selectivity between hFAAH

and rFAAH, this structure is simply an approximation of hFAAH. Humanized rFAAH is not

identical to hFAAH despite containing identical amino acid sequence within the active site.

Unknown secondary interactions within the tertiary structure of hFAAH must play a role in

the overall structure and complete substrate and inhibitor interactions that distinguish

human from rat FAAH. Differences in secondary structure due to alternative amino acids will have some effect, however small, on the inhibitor potencies. Otherwise, h/rFAAH would exhibit identical properties to hFAAH both in catalytic efficiency and inhibitor sensitivity profiles.

5.4 Discovery of Lead FAAH Inhibitors from Prostanoid Intermediates

Within the confines of our prostamide study, all intermediates from the extensive synthesis were subjected to biological testing in our laboratory to determine their activity on

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the cannabinoid receptors, CB1 and CB2, and the endocannabinoid hydrolyzing enzymes,

FAAH and MGL.† The assays were performed simply as a means to increase the library of compounds screened against these cannabinergic targets, and it was assumed that there would be no meaningful activity. This was especially true due to the high level of chirality and rigidity incorporated into our intermediates as it would be unusual for our compounds to exhibit the required spatial arrangement to bind an enzyme such as FAAH. It was surprisingly found that three of our intermediates, the enone and the S and R allylic alcohol

compounds, did, in fact, exhibit significant inhibition of FAAH. (Figure 5.14) Initial 3-point

screening resulted in 43% inhibition of rFAAH by the enone AM7609, 62% inhibition by R-

allylic alcohol AM7610, and 91% inhibition by S-allylic alcohol AM7611 all at 1 µM. With the

allylic alcohols exhibiting potency in approximately the nanomolar range, 8-point assays

were carried out and their curves are illustrated in Figure 5.15. AM7610 exhibited

inhibition with an IC50 of 3.9 µM while AM7611 demonstrated an IC50 of 800 nM.

Figure 5.14 Prostamide intermediates discovered to be lead FAAH inhibitor compounds

† The CB1, CB2, FAAH and MGL screening was done by Erin Shelnut, Kyle Whitten, Girija Rajarshi, and Pusheng Fan with assistance from Jodi Wood and Mahmoud Nasr

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Figure 5.15 Enzyme inhibition curves of R-allylic alcohol AM7610 and S-allylic alcohol AM7611

The 8-point assays resulted in IC50 values slightly greater than expected. However

compared to the affinity of the initial prototypes for the current generation of FAAH

inhibitors, both AM7610 and AM7611 are markedly potent. This set of bicyclic lactones

represents a promising foundation from which to optimize FAAH inhibition employing SAR

study. The potency increase upon conversion of the enone to the allylic alcohol and the

distinction in affinity between the R and S allylic alcohol stereoisomers, leads to the

intriguing possibility that hydrogen bonding of the 15-hydroxyl group to the enzyme plays

an important role in the inhibition. It also gives the impression that these compounds are

indeed inhibiting through a direct binding interaction with the enzyme.

Of the three intermediates, AM7609 and AM7610 additionally exhibited 50% inhibition of MGL at a concentration of about 100 µM based on 3-point assays. AM7611, the most potent of the rFAAH inhibitors, inhibited MGL by 50% in the range of 10-100 µM. Thus, although these compounds are able to bind MGL at elevated concentration, there is substantial selectivity for FAAH. When screened against hFAAH, none of the intermediates were able to inhibit this orthologue up to 100µM.

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Figure 5.16 Preliminary analogs in the identification of the prostamide intermediate pharmacophore for FAAH inhibition. Cleavage of the benzoate and of the terminal TBS groups eliminates activity.

With the discovery of these intermediates as relatively potent FAAH inhibitors, a further investigation into these chiral, bicyclic inhibitors was undertaken. Preliminary analogs were quickly synthesized wherein the removal of the TBS protection from the tail

20-hydroxyl as well as the removal the benzoyl protection from the 11-hydroxyl were employed to determine their effect on astivity. (Figure 5.16) This is the first step in

establishing a pharmacophore for our new class of FAAH inhibitors. Clearly, it is unusual for

a bioactive molecule to maintain a TBS protective group, and this moiety would rarely be

incorporated into the design of a therapeutic agent. In this case, it may simply serves as a

bulky, hydrophobic substituent at the tail position.

The results of 3-point screening on AM7648, AM7649, AM7650, AM7612 and AM7613

gave no indication of inhibitory activity up to 100 µM. Thus, because the polar free hydroxyl

eliminates activity, a bulky hydrophobic group is required at the tail position of the long

chain. Additionally, the benzoyl protective group is also required. It is unclear at this point,

whether the benzoyl group is similarly required to mask the polarity of the 11-hydroxyl or

whether this ester is required in order to interact with Ser241 in the active site of FAAH.

Further SAR analysis will provide more evidence as to which is the case.

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5.5 Design and Synthesis of Prostanoid Intermediate Analogs for FAAH Inhibition

By comparing our lead compounds with current FAAH inhibitors and understanding the mechanism by which they inhibit FAAH, we can postulate how our intermediates are interacting with FAAH to inhibit its actions. We hypothesize that the long lipophilic tail positions itself within the hydrophobic channel of the FAAH active site. This same channel is the locale for binding of the acyl arachidonyl portion of anandamide and the phenhexyl moiety of reversible inhibitors such as OL-135. Additionally, we can propose that one of the two ester moieties is able to interact with Ser241. It is uncertain whether the benzoate ester or the γ-butyrolactone is active towards nucleophilic attack by the Ser-Ser-Lys catalytic triad. The benzoate was determined to be essential for binding activity, and due to the electronegativity of the phenyl group, the carbonyl carbon of the benzoate is more activated towards nucleophilic attack. Both of these observations support the suggestion that the benzoate carbonyl is responsible for FAAH inhibition. However, it may be that the benzoate serves simply to convert the polar 11-hydroxyl into a more lipophilic group, and

that the chiral orientation of the γ-butyrolactone places it advantageously in line for bonding

the Ser241. A molecular modeling study would be extremely valuable for clarification of these hypotheses.

Figure 5.17 Representation of the proposed modifications to the lead compounds including increasing and decreasing the chain length and changing the benzoyl and TBS functionalities.

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An initial set of compounds were designed to assess the structure-activity

relationship of the inhibitors to rFAAH. The first round of analogs was set to include

modifications at the 11-hydroxyl group, at the 20-hydroxyl group, and of the chain length

between the 15-hydroxyl and ω tail. (Figure 5.17) Substituting the benzoyl group and/or

TBS group with groups of varying sizes, electronegativity, and lipophilicity will lead to a

better understanding of the requirements for binding. Additionally, changing the distance

between these moieties could lead us to discover the ideal placement of these essential

groups and give a better overall “fit” of the inhibitor to the enzyme. The first set of analogs

was chosen due to the ease of the synthesis from the steps previously established within

the prostamide synthesis. (Figure 5.18)

Figure 5.18 The first set of analogs synthesized for the study of FAAH inhibition.

The synthesis of the analogs in Figure 5.18 begins with the establishment of the phosphonates to be used in the Horner-Wadsworth-Emmons (HWE) reaction. Employing different chain lengths and using both straight carbon chains and those with hydroxyl functionality at the tail, various analogs can be easily reached. Similar to prostamide synthesis, methyl dimethyl phosphonate reacts with n-BuLi to form a phospho-methyl

-226- carbanion, and either a lactone of variable size or a methyl ester of variable size is added to make the desired phosphonate. (Scheme 5.1) The phosphonates containing the hydroxyl

Scheme 5.1 Establishment of the phosphonates

tail moiety are protected with TBS prior to the next step. A HWE reaction establishes the E- enone selectively, and the enone is then nonstereoselectively reduced to the R and S allylic alcohols. (Scheme 5.2) The stereoisomers are separated via flash chromatography, and at this point, the syntheses for compounds AM7656, AM7657, AM7658, AM7659 are complete.

For further functionalization of the tail group, the 8-carbon chain enone with a protected

Scheme 5.2 Horner-Wadsworth-Emmons and reduction to give AM7658, AM7659, AM7656, and AM7657

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hydroxyl tail is further derivatized. (Scheme 5.3) Following the HWE, the tail hydroxyl is deprotected and subsequently esterified to produce the acetate, the benzoate, the trimethylacetate, and the 1-adamantanecarborxylate using their respective acid chlorides.

The final step is to reduce the enone to the R and S allylic alcohols, and upon separation by flash chromatography, AM7660, AM7661, AM7662, AM7663, AM7664, AM7665, AM7666, and AM7667, are obtained. These syntheses could readily be extended further to provide numerous analogs, however the scope of this study was kept limited and finite.

Scheme 5.3 Horner-Wadsworth-Emmons, cleavage of the TBS protecting group, ester synthesis, and reduction to give AM7660, AM7661, AM7662, AM7663, AM7664, AM7665, AM7666, and AM7667.

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5.6 Evaluation of the Prostanoid Intermediate Analogs as FAAH Inhibitors

Table 5.2 Results of the 3pt screening of the prostamide intermediate FAAH inhibitor analogs

AM # Description 3pt rFAAH uM Estimated 3pt hFAAH IC50

7609 Bz-enone-OTBS 43% at 1µM* ≈1µM >100µM

7610 Bz-15R-OTBS 62% at 1µM* ≈1µM >100µM

7611 Bz-15S-OTBS 91% at 1µM* ≈100nM >100µM

7656 Bz-15S-(6C)-OTBS 9% at 1µM 10-100µM >100µM

7657 Bz-15R-(6C)-OTBS 18% at 1µM ≈10µM >100µM

7658 Bz-15S-12C 81% at 1µM* ≈100nM >100µM

7659 Bz-15R-12C 45% at 1µM* ≈1µM >100µM

7660 Bz-15S-OAc 18% at 1µM 1-10µM >100µM

7661 Bz-15R-OAc 6% at 1µM 10-100µM >100µM

7662 Bz-15S-OBz 72% at 1µM* <1µM >100µM

7663 Bz-15R-OBz 36% at 1µM 1-10µM >100µM

7664 Bz-15S-OAcMe3 81% at 1µM* <1µM >100µM

7665 Bz-15R-OAcMe3 42% at 1µM* ≈1µM >100µM 7666 Bz-15S-OAcAdm 90% at 1µM* ≈100nM >100µM

7667 Bz-15R-OAcAdm 67% at 1µM* <1µM >100µM

The results of the three point screening of this primary set of compounds are

tabulated in Table 5.2. The most evident finding is the consistency with which the S stereoisomers AM7611, AM7658, AM7660, AM7662, AM7664, and AM7666 show an

enhanced affinity over the R stereoisomers AM7610, AM7659, AM7661, AM7663, AM7665,

and AM7667. In every pairing of isomer analogs, the S stereoisomer shows a greater

potency for FAAH inhibition. Overall, using 1 µM the R isomers gave approximately half the

amount of inhibition the S isomers gave. This gives a strong indication that the hydroxyl is

involved in binding the active site and suggests that perhaps the 15-hydroxyl is

participating in a hydrogen bond interaction with a residue within the active site.

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Additionally, the orientation of the S isomer positions the hydroxyl in a more optimal

position within the binding pocket for either accepting or donating a hydrogen bond to a

nearby amino acid side chain.

Shortening the hydrocarbon chain between the main cyclopentyl ring and the bulky

tail OTBS group substantially decreases the affinity for FAAH. Compounds AM7656 and

AM7657, both containing only 6 carbons in their chain compared to the lead compounds’ 8 carbons, exhibit Kis in the 10-100 µM range. The increased length of the 12 carbon

nonfunctionalized chain analogs AM7658 and AM7659 are more accepted and give inhibition

very similar to that of the lead compounds.

The ester functionalized analogs reveal a trend that larger groups give greater

activity. The acetate AM7660, AM7661 exhibited the lowest inhibition at 1 µM, while

benzoate AM7662, AM7663 gave substantially greater inhibition, and trimethyl acetate

AM7664, AM7665 gave even slightly greater inhibition. The 1-adamantane carboxylate

compounds AM7666, AM7667, gave the greatest inhibition of the analogs, and its large bulk

and hydrophobicity suggests that the binding pocket for the lipophilic portion of the

inhibitors is in fact quite large.

With the selected collection of analogs synthesized in this study, inhibition did not

exceed that of the lead compounds. However, insight into the size requirements of the terminal group and the preferred distance between the main ring and the tail group for our compounds were gained. In summary, a hydrophobic chain longer that 6 carbons with a considerably bulky tail is the current working model for successful FAAH inhibitors with our bicyclic structure. The 15S hydroxyl is preferred over the 15R and enone due most likely to

a hydrogen bonding interaction with a residue within the active site of enzyme.

There is no discernible inhibition of hFAAH by any of the analogs tested, however

this does not rule out their usefulness as inhibitors. Overall, there is a dearth of information

surrounding the differences between rat and human FAAH inhibition, and development of

compounds selective for both rFAAH and hFAAH would do much to further our

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understanding of the structural requirement for human FAAH inhibition. All in vivo pharmalogical testing is done in animal models prior to being subjected to human trials.

Thus, a better understanding of the comparative profiles of rFAAH and hFAAH would eventually lead to compounds that are more efficacious in humans. As with many potential drugs, positive results of compounds that test well in animal studies do not necessarily extend to human trials. This is especially true in the case of FAAH inhibition, where disparity between the orthologs in vitro give a significant indication that positive results in murine in vivo studies will not necessitate success in humans. For example, Pfizer’s most promising

FAAH inhibitor PF-04456845 showed excellent pharmacokinetic properties in rats, mice, and dogs and was extremely effective against pain in rats, however when entered into human

Phase 2 clinical trials was found ineffective against osteoarthritis.56

We do not claim our compounds to be as effective at FAAH inhibition as the current

FAAH inhibitors, only that we have identified a new scaffold capable of inhibition of FAAH.

Further SAR studies may indeed lead to a compound with significant inhibitor effects. It is

challenging to compare the inhibition results of our compounds with those in the literature.

As mentioned earlier, studies giving “FAAH inhibition” results use rFAAH by default, and it is

unknown whether the inhibitors in literature are effective against hFAAH. If our results were

evaluated in a similar way, our inhibitors would be considered quite effective lead

compounds for FAAH. However, in light of the more recent establishment of substantial

binding differences between rFAAH and hFAAH, we postulate that our compounds will be

ineffective in the human orthologue.

Our compounds could lead to a better understanding of the differences between rat

and human FAAH active sites/selectivity. Upon examination of literature, it is difficult to

assess how many of the FAAH inhibitors claimed to date are indeed selective for rFAAH. Few

studies, up until recently, give the activity of inhibitors on hFAAH in addition to the rFAAH

data for comparison. It is possible that some of those compounds are selective for rFAAH.

However, it is also possible that our compounds are some of the only compounds that do

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not inhibit human FAAH to any measurable extent. They are undeniably selective for rFAAH

over hFAAH, and until all of the known FAAH inhibitors are tested against hFAAH the

uniqueness of our rFAAH selective inhibitors is unknown.

5.7 Future Directions for FAAH Inhibition by Prostanoid Intermediates

There are several areas in which the FAAH inhibitor project could be expanded. This study is just the primary investigation from which a comprehensive design of experiments

(DOE) could be implemented. Before expanding the library of compounds, a number of analyses of the current lead compounds could be performed to create a foundation for improvement. Molecular modeling of our compounds within the active site of rFAAH and the humanized rFAAH crystal structures would give invaluable information about their binding mode. Biochemical assays could give indications of the nature of inhibitory interaction between the inhibitor and the enzymes. Based on the results of these studies, further inhibitors can be designed and synthesized for additional SAR analysis and optimization.

5.7.1 Molecular Modeling for Exploration of the Binding Site Interactions

Molecular modeling studies would allow us to take advantage of the established structural biology of the FAAH enzymes and design a new set of inhibitors optimized towards binding the active site. Overlaying the structures of our compounds with those of the bound substrate and recent inhibitors, would give better understanding of the orientation of our intermediates and how they fulfill the requirements for binding. Within this study, we have begun probing the large lipophilic pocket to see how large a group would be tolerated. However, analysis of the docking of our structures may give better insight into how expandable this pocket is and what size group would be optimal for inhibition. Additionally, comparison of the positions of the R- and S-15 hydroxyl groups when bound would allow us to determine which, if any, of the residues are within

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reasonable range for hydrogen bonding. We can also gain a better perspective on which of

the two ester groups, if either, are responsible for interaction with the Ser-Ser-Lys catalytic

triad from their orientation and proximity. Contrasting the molecular docking of these

inhibitors to rFAAH and to h/rFAAH would lead to a more advanced understanding of their

selectivity for rFAAH over hFAAH. Identifying which interactions are favorable to binding

rFAAH that are not present in binding h/rFAAH would allow for the design of compounds

more effective against hFAAH. Thus, a whole host of valuable information can be gained

from molecular modeling studies to help further the understanding of how compounds bind

and to aid in the design of more potent inhibitors.

5.7.2 Biochemical Assays to Assess Reversibility, Competitiveness, and Selectivity

Biochemical assays could give valuable insight into how our intermediates are

functioning to inhibit FAAH. Determination of whether the inhibition by these intermediates

is reversible or irreversible can be easily ascertained by incubating the inhibitors with the

enzymes for longer periods of time to see if their efficacy is enhanced. Unlike reversible

inhibitors, which reach equilibrium between the bound and unbound states, irreversible

inhibitors continue to bind the enzyme until there is a saturation of inhibitors bound to the

enzymes. Thus, the longer inhibitors are allowed to act on the enzymes, the greater the

amount of inhibitor-bound enzymes and the higher the observed potency. This explains why the potency of irreversible inhibitors cannot correctly be expressed as an IC50 value but

instead should be written as Kinact/Ki. With these values, greater Kinact/Ki numbers denote a higher potency. Thus, if the bioassay shows an equilibration of inhibitory action where the

IC50 remains constant, the compound is said to be reversible, and if longer incubation leads

to increased potency, it is irreversible. Alternatively, dialisis experiments can be run to see if

the inhibitor is separable from the enzyme by passing through a membrane. If the

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compound is nondializable, there is an irreversible bond formed between the inhibitor and

the enzyme.

A

B C

Figure 5.19 The kinetic curves of substrate binding enzymes. (A) Comparison between inhibition of the rate of enzymatic reaction with increasing substrate concentration by a competitive inhibitor and by a noncompetitive inhibitor. (B) The Lineweaver-Burke (double reciprocal) plot of increasing levels of competitive inhibitor highlighting the identical VMAX and different KM values. (C) The Lineweaver- Burke (double reciprocal) plot of increasing levels of noncompetitive inhibitor highlighting the different VMAX and identical KM values.

Through analysis of biological assays, we can also determine whether our inhibitors

are competitive or noncompetitive. Several of the current known FAAH inhibitors are, in

fact, noncompetitive inhibitors; meaning that they do not inhibit by binding directly to the

active site but instead bind in a disparate location where they employ conformational

changes to inhibit binding of the substrate. Thus they do not compete directly with the substrate for binding of the active site. Curves of the enzyme kinetics prove useful in determining whether an inhibitor is competitive or noncompetitive. In competitive inhibition, because the substrate and enzyme are acting on the same site, the substrate concentration can be increased to overcome the effects of the inhibitor and saturate the enzyme. Thus, in terms of the Lineweaver-Burke double reciprocal interpretation, the Vmax, which is a

-234- measure of reaction rate, will remain the same regardless of the inhibitor concentration.

(Figure 5.19) However, it will require a higher concentration of substrate to overcome occupation by the inhibitor and the Km, which is a measure of inhibitor potency, will therefore be higher. Conversely, in noncompetitive inhibition, the substrate and enzyme are not competing with each other to bind the active site, and increasing substrate concentration cannot overcome the inhibition. In this case, the Vmax will be reduced by increasing levels of inhibitor and the Km will remain unchanged. If our compounds were found to be noncompetitive inhibitors, the success of their inhibition would not necessarily be due to interactions with the Ser-Ser-Lys catalytic triad. The inhibitors could instead be binding elsewhere and inciting a conformational change in the active site that renders it inactive to enzymatic conversion of the substrate.

Finally, bioassays screening our inhibitors against several enzymes within the serine hydrolase class would facilitate the determination of the overall selectivity for FAAH. While it is uncertain whether these intermediates selectively recognize FAAH as a target for inhibition, we can postulate that the rigidity and chirality of our structures may enhance their selectivity. Frequently, employing more constrained structural features and incorporating chiral moieties optimized to the structural biology of the enzyme will allow the inhibitor to “fit” preferentially into the target enzyme and be sterically unable to bind the others.

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5.7.3 Synthetic Routes for an Extended Series of Prostanoid Intermediate Test Compounds

Figure 5.20 Proposed synthesis for the enone intermediate of future analogs

To further the FAAH inhibitor project, the library of test compounds can very readily

be expanded to include a large number of analogs. Few synthetic steps are needed to synthesize a diversity of analogs, and the methodology involved is ideal for parallel synthesis. The methods already established in the synthesis of the current library could be extended to create longer hydrocarbon chains by employing longer, functionalized phosphonates. (Figure 5.20) A ginormous number of modifications can be made to the tail

alcohol including ethers, carbamates, esters imines, secondary and tertiary amines, and

enol thioesters all with the ability to incorporate differing R groups including a significant

number of known biologically active heterocyclic groups. Once the primary alcohol is deprotected it can be transformed into a leaving group and upon treatment with base give an alkene. (Figure 5.21) This alkene can then be functionalized to several different groups

by adding across the double bond including, but not limited to, ethers by treatment with

alcohols and a cyano by treatment with HCN. Additionally, treating the free alcohol with an

isocyanate leads to the formation of a carbamate. (Figure 5.22) This functionality could be

of interest due to the known preference of FAAH for carbamates. Alternatively, the tail

alcohol can be oxidized to an aldehyde giving way to even more possible functionalities.

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Figure 5.21 Proposed synthesis for the establishment of future ether and cyano analogs

Figure 5.22 Proposed synthesis for the establishment of carbamates and of future ester, imine, secondary amine, tertiary amine, and enol thioether analogs through an initial conversion to aldehyde.

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Esters are formed upon treatment with alcohols, imines upon treatment with aryl amines,

secondary amines upon treatment with primary amines and hydrogenation catalyst (and

also tertiary amines from secondary amines), and enol thioesters upon treatment with thiols

and titanium tetrachloride. The terminal alcohol could be oxidized further to a carboxylic

acid which allows for yet more synthetic possibilities. Most of these functional changes can also be applied to the 11-hydroxyl moiety. Following cleavage of the benzoyl group, the free

alcohol can then be acted on in a similar manner presented here for the tail hydroxyl. The

reactions may tend to be a little slower due to the secondary nature of the 11-hydroxyl.

The (+)-Corey lactone benzoate aldehyde, with absolute inversion in stereochemistry

to the standard prostaglandin synthesis starting material, is commercially available. It would

be worthwhile to synthesize a few representative compounds with this inverted

stereochemistry and test its inhibitory ability. Comparison of their affinities to that of an

analog containing identical functionality but opposite chirality would lead to a determination

of the importance of chirality. It may support or disprove our proposal that the

stereochemistry of the core bicyclic structure is integral for binding rFAAH. The synthesis

would be identical to that of the corresponding entantiomer analog. Thus, there is a broad

list of analogs that can be synthesized in relatively short order to test in further

investigation into the SAR of FAAH.

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Appendix 1: Supplemental Synthetic Methods and Characterization for Compounds Presented in Chapter 2

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General Information:

All reactions were carried out in flame-dried glassware under argon atmosphere. Corey’s lactone benzoate aldehyde starting material was purchased from Cayman Chemical and used as supplied. All other reagents were purchased from Sigma-Aldrich and used as supplied. Tetrahydrofuran, dichloromethane, diethyl ether, and acetonitrile were obtained from a dry solvent system (alumina) and used without further drying. Unless otherwise noted, reactions were magnetically stirred and monitored by thin layer chromatography with Merck 250 µm silica gel 60 Å plates. Flash chromatography was performed using Biotage Isolera Flash Purification with Luknova 40-60 µm 60 Å silica gel columns. Yields refer to chromatographically and spectroscopically pure compounds, unless otherwise noted. 1H spectra were taken in CDCl3, at 400 or 500 MHz as indicated. 13C NMR spectra were also taken in CDCl3 at 100 MHz. Chemical shifts are reported in parts per million relative to TMS 1 13 ( H, δ 0.00) or CDCl3 ( C, δ 77.0). Data are reported as follows: chemical shift, multiplicity (s = singlet, d = doublet, t = triplet, q = quartet, quin = quintet, m = multiplet, br = broad), coupling constant, integration. Diastereomeric ratios were determined by 1H NMR (500 MHz) analysis of crude mixtures. High resolution mass- spectra were obtained on a Waters Q-TOF Ultima spectrometer at University of Illinois, SCS, Mass Spectrometry Laboratory.

Experimental Methods

Preparation of Dimethyl 7-hydroxy-2-oxoheptylphosphonate

O MeO P OMe

OH O A solution of dimethyl methylphosphonate (3.0 mL, 28.1 mmol) in anhydrous THF (90 mL) was cooled to -78 ºC under argon. n-BuLi (2.5 M in hexane, 12.0 mL, 30.0 mmol) was added dropwise and stirring was continued at -78 ºC for 90 min. ε- caprolactone (1.5 mL, 13.5 mmol) was added and after 1 hr, oxalic acid (2.60g, 28.8mmol) was added and stirring was continued for 30 min warming to rt. The reaction mixture was filtered over Celite, washed with THF, and concentrated. Purification by flash chromatography on silica gel (MeOH/DCM, 5:95) afforded 3.16g 1 (98%) of the desired product as a colorless oil. H NMR (400MHz, CDCl3) δ [ppm] = 3.79 (d, JHP = 11.54 Hz, 6H), 3.62 (t, J = 6.53 Hz, 2H), 3.10 (d, JHP = 23.09 Hz, 2H), 2.64 (t, J = 7.28 Hz, 2H), 2.47 (br. s., 1H) 1.66-1.53 (m, 4H), 1.41 – 1.35 (m, 2H). + HRMS calcd for C9H20O5P, [M+H ] 239.1048, found 239.1043.

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Dimethyl 7-(tert-butyldimethylsilyloxy)-2-oxoheptylphosphonate (2)

O MeO P OMe

OTBS O To a solution of dimethyl 7-hydroxy-2-oxoheptylphosphonate (2.31 g, 9.68 mmol) in anhydrous DCM (60 mL) at 0ºC was added imidazole (1.36 g, 20.35 mmol) and tert-butylchlorodimethylsilane (3.03 g, 20.10 mmol). The reaction was allowed to stir overnight and slowly warm to rt. The mixture was quenched with saturated NH4Cl solution. The organic phase was washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (Acetone/Hexanes, 80:20) afforded 3.34g (98%) of the desired product as a 1 colorless oil. H NMR (400MHz, CDCl3) δ [ppm] = 3.79 (d, JHP = 11.54 Hz, 6H), 3.60 (t, J = 6.53 Hz, 2H), 3.09 (d, JHP = 23.09 Hz, 2H), 2.62 (t, J = 7.28 Hz, 2H), 1.60 (quin, J = 7.53 Hz, 2H), 1.52 (quin, J = 7.03 Hz, 2H), 1.37 – 1.29 (m, 2H), 0.89 (s, + 9H), 0.04 (s, 6H). HRMS calcd for C15H34O5SiP, [M+H ] 353.1913, found 353.1905.

(3aR,4R,5R,6aS)-4-((E)-8-(tert-butyldimethylsilyloxy)-3-oxooct-1-enyl-2- oxohexahydro-2H-cyclopenta[b]furan-5-yl benzoate (3)

O O

O O OTBS O

To a solution of NaH (60% dispersion in mineral oil, 112.5 mg, 2.82 mmol) in anhydrous THF (120 mL) at 0 ºC was added phosphonate 2 (892.1 mg, 2.53 mmol). The reaction was stirred at rt for 1hr after which it was cooled to 0 ºC and corey’s lactone benzoate aldehyde (696.3 mg, 2.539 mmol) was added. After 2.5 hrs at rt the reaction was quenched with glacial acetic acid (0.5 mL). The mixture was concentrated, diluted with EtOAc, and washed with water and brine. The organic layers were combined, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 90:10) afforded 1.27g (100%) of the 1 desired product as a colorless oil. H NMR (400MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.20 Hz, 2H), 7.58 (t, J = 7.54 Hz, 1H), 7.45 (t, J = 7.54 Hz, 2H), 6.70 (dd, J = 15.77 Hz, 7.54 Hz, 1H), 6.24 (d, J = 15.77 Hz, 1H), 5.33 (q, J = 5.49 Hz, 1H), 5.11 (t, J = 4.80 Hz, 1H), 3.59 (t, J = 6.34 Hz, 2H), 2.97 – 2.86 (m, 3H), 2.63 (dt, J = 15.77 Hz, 6.51 Hz, 1H), 2.58 – 2.47 (m, 3H), 2.30 (ddd, J = 15.63, 3.91, 2.00 Hz, 1H), 1.61 (quin, J = 7.54 Hz, 2H), 1.52 (quin, J = 6.86 Hz, 2H), 1.39 – 1.30 (m, + 2H), 0.89 (s, 9H), 0.04, (s, 6H). HRMS calcd for C28H41O6Si, [M+H ] 501.2672, found 501.2677.

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(3aR,4R,5R,6aS)-4-((R/S,E)-8-(tert-butyldimethylsilyloxy)-3-hydroxyoct-1- enyl)-2-oxohexhydro-2H-cyclopenta[b]furan-5-yl benzoate (4)

O O O O

O O O OTBS O OTBS OH OH

A solution of enone 3 (1.27 g, 2.54 mmol) in MeOH (100mL) was cooled to 0 ºC. Cerium (III) chloride heptahydrate (1.418 g, 3.81mmol) and sodium borohydride (144 mg, 3.81mmol) were added sequentially and the mixture was stirred for 30 min. It was concentrated, diluted with EtOAc, washed with water and brine, dried over MgSO4, and concentrated a second time. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 60:40) gave pure 15S and 15R isomers in 10% de for a total of 1.09 g (86%) [0.60 g (S-isomer) and 0.49 g (R-isomer)] as a colorless oil. 1 Isomer S H NMR (400 MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.32 Hz, 2H), 7.57 (t, J = 7.32 Hz, 1H), 7.44 (t, J = 7.81 Hz, 2H), 5.65 (dd, J = 15.63 Hz, 6.10 Hz, 1H), 5.56 (dd, J = 15.38 Hz, 7.08 Hz, 1H), 5.23 (q, J = 5.86 Hz, 1H), 5.06 (t, J = 5.62 Hz, 1H), 4.08 (q, J = 6.10 Hz, 1H), 3.56 (t, J = 6.47 Hz, 2H), 2.91 – 2.77 (m, 2H), 2.74 (q, J = 6.35 Hz, 1H), 2.61 (dt, J = 15.38 Hz, 6.59 Hz, 1H), 2.52 (d, J = 16.6 Hz, 1H), 2.23 (dd, J = 15.38 Hz, 3.91 Hz, 1H), 1.70 (br. s., 1H) 1.55 – 1.40 (m, 4H), + 1.40 – 1.25 (m, 4H), 0.89 (s, 9H), 0.03 (s, 6H). HRMS calcd for C28H43O6Si, [M+H ] 1 503.2829, found 503.2816. Isomer R H NMR (400 MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.32 Hz, 2H), 7.56 (t, J = 7.32 Hz, 1H), 7.44 (t, J = 7.81 Hz, 2H), 5.66 (dd, J = 15.63 Hz, 5.62 Hz, 1H), 5.58 (dd, J = 15.38 Hz, 6.84 Hz, 1H), 5.24 (q, J = 5.62 Hz, 1H), 5.06 (t, J = 5.62 Hz, 1H), 4.09 (q, J = 5.86 Hz, 1H), 3.58 (t, J = 6.35 Hz, 2H), 2.91 – 2.78 (m, 2H), 2.74 (q, J = 6.35 Hz, 1H), 2.60 (dt, J = 15.38 Hz, 6.59 Hz, 1H), 2.51 (d, J = 16.6 Hz, 1H), 2.23 (dd, J = 15.38 Hz, 3.91 Hz, 1H), 1.77 (br. s., 1H) 1.55 – 1.45 (m, 4H), 1.40 – 1.25 (m, 4H), 0.89 (s, 9H), 0.04 (s, 6H). HRMS + calcd for C28H43O6Si, [M+H ] 503.2829, found 503.2826.

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General procedure for mosher ester synthesis

O O O O S Mosher of 15S-enol S Mosher of 15R-enol O O O OTBS O OTBS O O O O

CF3 CF3 Ph OMe Ph OMe

O O O O R Mosher of 15S-enol R Mosher of 15R-enol O O O OTBS O OTBS O O O O

CF3 CF3 MeO Ph MeO Ph To a solution of (+) or (-) enol 4 (12.0 mg, 0.024 mmol) in anhydrous DCM (2mL) was added anhydrous pyridine (10 µL, 0.124 mmol) followed by the addition of the respective mosher acid chloride (R-(-)-MTPA-Cl or S-(+)-MTPA-Cl) (14 µL, 0.073 mmol). The reaction was stirred at rt for 2 hrs at which point it was diluted with DCM and washed with water and brine. The organic layers were combined, dried over MgSO4, filtered and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 40:60) afforded the desired product as a colorless oil (82- 94%).

1 S Mosher of 15S Allylic Alcohol H NMR (500 MHz, CDCl3) δ [ppm] = 7.98 (d, J = 7.20 Hz, 2H), 7.57 (t, J = 7.54 Hz, 1H), 7.52 – 7.47 (m, 2H), 7.47 – 7.36 (m, 5H), 5.71 (dd, J = 15.43 Hz, 7.20 Hz, 1H), 5.60 (dd, J = 15.43 Hz, 7.54 Hz, 1H), 5.40 (q, J = 7.20 Hz, 1H) 5.22 (q, J = 6.17 Hz, 1H), 5.03 (t, J = 5.31 Hz, 1H), 3.51 – 3.47 (m, 5H), 2.87 – 2.69 (m, 3H), 2.58 (dt, J = 15.43 Hz, 6.86 Hz, 1H), 2.42 (d, J = 17.49 Hz, 1H), 2.22 (dd, J = 15.43 Hz, 5.49 Hz, 1H) 1.72 – 1.51 (m, 2H), 1.35 (quin, J = 6.43 Hz, 2H), 1.27 – 1.12 (m, 4H), 0.88 (s, 9H), 0.03 (s, 6H)

1 S Mosher of 15R Allylic Alcohol H NMR (500 MHz, CDCl3) δ [ppm] = 7.97 (d, J = 7.55 Hz, 2H), 7.56 (t, J = 7.20 Hz, 1H), 7.52 – 7.46 (m, 2H), 7.43 (t, J = 7.89 Hz, 2H), 7.40 – 7.35 (m, 3H), 5.60 – 5.50 (m, 2H), 5.41 (q, J = 5.83 Hz, 1H), 5.16 (q, J = 5.14 Hz, 1H), 5.03 (t, J = 6.17 Hz, 1H), 3.56 (t, J = 6.51 Hz, 2H), 3.48 (s, 3H), 2.84 (dd, J = 18.17 Hz, 9.94 Hz, 1H), 2.76 – 2.65 (m, 2H), 2.50 (dt, J = 15.43Hz, 6.51Hz, 1H), 2.45 (d, J = 18.17 Hz, 1H), 2.22 (dd, J = 15.43 Hz, 3.43 Hz, 1H), 1.80 – 1.69 (m, 1H), 1.68 – 1.58 (m, 1H), 1.50 – 1.43 (m, 2H), 1.37 – 1.28 (m, 4H), 0.89 (s, 9H), 0.04 (s, 6H).

1 R Mosher of Allylic Alcohol H NMR (500 MHz, CDCl3) δ [ppm] = 7.97 (d, J = 7.20 Hz, 2H), 7.56 (t, J = 7.54 Hz, 1H), 7.52 – 7.47 (m, 2H), 7.47 – 7.36 (m, 5H), 5.61 (dd, J = 15.43 Hz, 7.54 Hz, 1H), 5.49 (dd, J = 15.09 Hz, 7.20 Hz, 1H), 5.39 (q, J = 6.86 Hz, 1H), 5.18 (q, J = 5.49 Hz, 1H), 5.02 (t, J = 6.53 Hz, 1H), 3.56 – 3.51 (m, 5H), 2.83 (dd, J = 17.83 Hz, 9.60 Hz, 1H), 2.78 – 2.65 (m, 2H), 2.53 (dt, J = 15.43 Hz, 6.51 Hz, 1H), 2.43 (d, J = 17.83 Hz, 1H), 2.20 (dd, J = 15.43 Hz, 4.11 Hz, 1H),

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1.75 – 1.54 (m, 2H), 1.45 – 1.36 (m, 2H), 1.33 – 1.20 (m, 4 H), 0.88 (s, 9H), 0.03 (s, 6H).

1 R Mosher of Allylic Alcohol H NMR (500 MHz, CDCl3) δ [ppm] = 7.97 (d, J = 7.20 Hz, 2H), 7.56 (t, J = 7.54 Hz, 1H), 7.51 – 7.46 (m, 2H), 7.43 (t, J = 7.89 Hz, 2H), 7.40 – 7.33 (m, 3H), 5.71 (dd, J = 15.43 Hz, 6.51 Hz, 1H) 5.64 (dd, J = 15.43 Hz, 6.51 Hz, 1H), 5.42 (q, J = 6.86 Hz, 1H), 5.21 (q, J = 5.49 Hz, 1H), 5.05 (t, J = 5.31 Hz, 1H), 3.54 (t, J = 6.51 Hz, 2H), 3.42 (s, 3H), 2.86 (dd, J = 17.83 Hz, 9.6 Hz, 1H), 2.80 – 2.72 (m, 2H), 2.56 (dt, J = 15.77 Hz, 6.51 Hz, 1H), 2.47 (d, J = 17.49 Hz, 1H), 2.23 (dd, J = 15.45 Hz, 3.43 Hz, 1H), 1.74 – 1.54 (m, 2H), 1.40 (quin, J = 6.52 Hz, 2H), 1.32 – 1.15 (m, 4H), 0.89 (s, 9H), 0.03 (s, 6H).

(3aR,4R,5R,6aS)-4-((S,E)-8-(tert-butyldimethylsilyloxy)-3-hydroxyoct-1- enyl)-5-hydroxyhexahydro-2H-cyclopenta[b]furan-2-one (5-S)

O O

HO OTBS OH To a solution of benzoate 4-S (955.1 mg, 1.90 mmol) in anhydrous THF (10mL) was added anhydrous MeOH (125 mL) followed by K2CO3 (262.6 mg, 1.90 mmol). The reaction was brought to 50 ºC and stirred for 2.5 hrs. Saturated NH4Cl (4mL) was added to quench the reaction and after 20 min the mixture was filtered through Celite, dried thoroughly over MgSO4, and concentrated. Purification by flash chromatography on silica gel (Acetone/DCM, 60:40) afforded 685.5 mg (91%) of the 1 desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.63 (dd, J = 15.56 Hz, 6.02 Hz, 1H), 5.50 (dd, J = 15.56 Hz, 7.53 Hz, 1H), 4.93 (td, J = 7.03 Hz, 3.01 Hz, 1H), 4.09 (q, J = 6.02 Hz, 1H), 3.99 (q, J = 7.03 Hz, 1H), 3.60 (t, J = 6.53 Hz, 2H), 2.92 (br. s., 1H), 2.75 (dd, J = 17.57 Hz, 9.54 Hz, 1H), 2.63 (q, J = 7.53 Hz, 1H), 2.51 – 2.42 (m, 2H), 2.34 (q, J = 7.53 Hz, 1H), 2.24 (br. s., 1H), 1.97 (ddd, J = 14.81 Hz, 6.78 Hz, 2.51 Hz, 1H), 1.58 – 1.47 (m, 4H), 1.44 – 1.28 (m, + 4H), 0.89 (s, 9H), 0.05 (s, 6H). HRMS calcd for C21H39O5Si, [M+H ] 399.2567, found 399.2564.

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(3aR,4R,5R,6aS)-4-((R,E)-8-(tert-butyldimethylsilyloxy)-3-hydroxyoct-1- enyl)-5-hydroxyhexahydro-2H-cyclopenta[b]furan-2-one (5-R)

O O

HO OTBS OH

To a solution of benzoate 4-R (786.4 mg, 1.56 mmol) in anhydrous THF (5 mL) was added anhydrous MeOH (60 mL) followed by K2CO3 (216.2 mg, 1.56 mmol). The reaction was brought to 50 ºC and stirred for 2.5 hrs. Saturated NH4Cl (2mL) was added to quench the reaction and after 20 min the mixture was filtered through Celite, dried thoroughly over MgSO4, and concentrated. Purification by flash chromatography on silica gel (Acetone/DCM, 60:40) afforded 553.2 mg (89%) of the 1 desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.57 (dd, J = 15.06 Hz, 7.03 Hz, 1H), 5.42 (dd, J = 15.56 Hz, 8.53 Hz, 1H), 4.89 (td, J = 7.03 Hz, 3.01 Hz, 1H), 4.02 (q, J = 6.86 Hz, 1H), 3.91 (q, J = 7.53 Hz, 1H), 3.91 (br. s., 1H), 3.60 (t, J = 6.53 Hz, 2H), 3.10 (br. s., 1H), 2.72 (dd, J = 18.07 Hz, 9.54 Hz, 1H), 2.60 – 2.48 (m, 2H), 2.40 (d, J = 18.07 Hz, 1H), 2.24 (d, J = 8.53 Hz, 1H), 1.91 (ddd, J = 14.56 Hz, 8.03 Hz, 3.01 Hz, 1H), 1.68 – 1.42 (m, 4H), 1.40 – 1.25 (m, + 4H), 0.89 (s, 9H), 0.05 (s, 6H). HRMS calcd for C21H39O5Si, [M+H ] 399.2567, found 399.2558.

(3aR,4R,5R,6aS)-4-((S,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-(triisopropylsilyloxy)hexahydro-2H- cyclopenta[b]furan-2-one (6-S)

O O

TIPSO OTBS OTIPS

A solution of diol 5-S (685.5 mg, 1.72 mmol) in anhydrous DCM (150mL) was cooled to 0 ºC and 2,6-lutidine (4.0mL, 34.45 mmol) was added. Upon addition of triisopropyl trifluoromethanesulfonate (4.6mL, 17.05 mmol) the reaction was stirred for 1 hr. It was quenched with saturated NH4Cl, extracted with DCM, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 10:90) afforded 1.099 g (90%) of the 1 desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.49 (dd, J = 15.63, 6.35 Hz, 1 H), 5.38 (dd, J=15.63, 7.81 Hz, 1 H) 5.02 (t, J=6.10 Hz, 1 H), 4.12 - 4.19 (m, 2 H), 3.59 (t, J=6.35 Hz, 2 H), 2.80 (dd, J=17.58, 10.74 Hz, 1 H), 2.70 - 2.76 (m, 1 H), 2.56 - 2.65 (m, 2 H), 2.17 (ddd, J=14.65, 6.59, 5.13 Hz, 1 H), 2.11 (d, J=14.65 Hz, 1 H), 1.53 - 1.42 (m, 4 H), 1.22 - 1.36 (m, 4 H), 1.05 (d, J=9.28 Hz, 42 H), 0.89 (s, 9 H), 0.04 (s, 6 H). HRMS calcd for C39H78O5NaSi3, [M+Na+] 733.5055, found 733.5055.

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(3aR,4R,5R,6aS)-4-((R,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-(triisopropylsilyloxy)hexahydro-2H- cyclopenta[b]furan-2-one (6-R)

O O

TIPSO OTBS OTIPS

A solution of diol 5-R (550.3 mg, 1.38 mmol) in anhydrous DCM (100 mL) was cooled to 0 ºC and 2,6-lutidine (3.2 mL, 27.8 mmol) was added. Upon addition of triisopropyl trifluoromethanesulfonate (3.73 mL, 13.82 mmol) the reaction was stirred for 1 hr. It was quenched with saturated NH4Cl, extracted with DCM, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 10:90) afforded 912.9 mg (93%) of 1 the desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.49 (dd, J=15.63, 6.84 Hz, 1 H), 5.36 (dd, J=15.63, 7.32 Hz, 1 H), 5.00 (t, J=6.35 Hz, 1 H), 4.17 (q, J=6.35 Hz, 1 H), 4.12 (q, J=4.07 Hz, 1 H), 3.59 (t, J=6.59 Hz, 2 H), 2.79 (dd, J=17.09, 10.74 Hz, 1 H), 2.69 - 2.76 (m, 1 H), 2.53 - 2.63 (m, 2 H), 2.22 (dt, J=14.65, 6.10 Hz, 1 H), 2.07 (d, J=15.14 Hz, 1 H), 1.43 - 1.61 (m, 4 H), 1.21 - 1.37 (m, 4 H), 1.04 (d, J=6.35 Hz, 42 H), 0.89 (s, 9 H), 0.06 (s, 6 H). HRMS calcd for + C39H79O5Si3, [M+H ] 711.5235, found 711.5233.

(+/-) (3aR,4R,5R,6aS)-4-((S,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-(triisopropylsilyloxy)hexahydro-2H- cyclopenta[b]furan-2-ol (7-S)

OH O

TIPSO OTBS OTIPS To a solution of lactone 6-S (1.099 g, 1.55 mmol) in anhydrous DCM at -78 ºC was added DIBAL-H (1.0 M in hexane, 2.78 mL, 2.78 mmol) dropwise. After 30 min the reaction mixture was quenched with saturated NH4Cl and gradually warmed to rt. It was diluted with DCM, and saturated sodium potassium tartrate was added. After stirring 20 min the phases were separated and the aqueous layer was backwashed twice with DCM. The organic layers were combined, dried on MgSO4, filtered, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 20:80) afforded 1.03 g (93%) of the desired product as a colorless 1 oil. H NMR (400 MHz, CDCl3) δ [ppm] (as a mixture of ~50:50 isomers – doubled integration) = 5.62 (br. s., 1 H), 5.41 - 5.53 (m, 5 H), 5.20 (d, J=9.60 Hz, 1 H), 4.63 - 4.75 (m, 2 H), 4.10 - 4.21 (m, 3 H), 4.03 (q, J=6.06 Hz, 1 H), 3.59 (t, J=6.52 Hz, 4 H), 2.85 (br. s., 1 H), 2.63 (br. s., 1 H), 2.49 - 2.60 (m, 2 H), 2.31 - 2.45 (m, 2 H), 2.16 - 2.29 (m, 3 H), 2.00 - 2.10 (m, 3 H), 1.75 - 1.85 (m, 1 H), 1.41 - 1.60 (m, 8 H), 1.21 - 1.37 (m, 8 H), 0.98 - 1.14 (m, 84 H), 0.89 (s, 18 H), 0.04 (s, 12 + H). HRMS calcd for C39H80O5NaSi3, [M+Na ] 735.5211, found 735.5232.

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(+/-) (3aR,4R,5R,6aS)-4-((R,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-(triisopropylsilyloxy)hexahydro-2H- cyclopenta[b]furan-2-ol (7-R)

OH O

TIPSO OTBS OTIPS

To a solution of lactone 6-R (912.9 mg, 1.28 mmol) in anhydrous DCM at -78 ºC was added DIBAL-H (1.0 M in hexane, 2.31 mL, 2.31 mmol) dropwise. After 30 min the reaction mixture was quenched with saturated NH4Cl and gradually warmed to rt. It was diluted with DCM, and saturated sodium potassium tartrate was added. After stirring 20 min the phases were separated and the aqueous layer was backwashed twice with DCM. The organic layers were combined, dried on MgSO4, filtered, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 20:80) afforded 849.0 mg (93%) of the desired product as a colorless oil. 1H NMR (400 MHz, CDCl3) δ [ppm] (as a mixture of ~50:50 isomers – doubled integration) = 5.49 (dd, J=15.63, 6.84 Hz, 1 H), 5.36 (dd, J=15.63, 7.32 Hz, 1 H), 5.00 (t, J=6.35 Hz, 1 H), 4.17 (q, J=6.35 Hz, 1 H), 4.12 (q, J=4.07 Hz, 1 H), 3.59 (t, J=6.59 Hz, 2 H), 2.79 (dd, J=17.09, 10.74 Hz, 1 H), 2.69 - 2.76 (m, 1 H), 2.53 - 2.63 (m, 2 H), 2.22 (dt, J=14.65, 6.10 Hz, 1 H), 2.07 (d, J=15.14 Hz, 1 H), 1.43 - 1.61 (m, 4 H), 1.21 - 1.37 (m, 4 H), 1.04 (d, J=6.35 Hz, 42 H), 0.89 (s, 9 H), 0.06 (s, 6 H). HRMS + calcd for C39H79O5Si3, [M+H ] 711.5235, found 711.5233.

(Z)-7-((1R,2R,3R,5S)-2-((S,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-hydroxy-3- (triisopropylsilyloxy)cyclopentyl)hept-5-enoic acid (8-S)

HO COOH

OTBS TIPSO OTIPS A suspension of (4-carboxybutyl)-triphenylphosphonium bromide (1.59 g, 3.59 mmol) in anhydrous THF (100 mL) was cooled to 0 ºC and potassium tert- butoxide (1.0 M in THF, 3.59 mL, 3.59 mmol) was added dropwise. The orange reaction mixture was stirred for 30 min at 0 ºC at which point lactol 7-S (512.7 mg, 0.719 mmol) in anhydrous THF (10mL) was added dropwise. The reaction continued for 1hr and was quenched with saturated NH4Cl and warmed to rt. After separation the organic layer was evaporated, diluted with EtOAc, washed with water and brine, dried over MgSO4, filtered, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 50:50) afforded 416.7 mg (73%) of the desired 1 product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.56 – 5.31 (m, 4H), 4.20 – 4.12 (m, 3H), 3.60 (t, J = 6.78 Hz, 2H), 2.45 – 2.32 (m, 4H), 2.25 – 2.10 (m, 3H), 1.99 (d, J = 14.05 Hz, 1H), 1.81 (dt, J = 13.93 Hz, 4.33 Hz, 1H), 1.71 (quin, J = 7.28 Hz, 2H), 1.62 – 1.42 (m, 5H), 1.36 – 1.23 (m, 4H), 1.08 – 1.02 (m,

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+ 42H), 0.89 (s, 9H), 0.05 (s, 6H). HRMS calcd for C44H88O6NaSi3, [M+Na ] 819.5786, found 819.5790.

(Z)-7-((1R,2R,3R,5S)-2-((R,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-hydroxy-3- (triisopropylsilyloxy)cyclopentyl)hept-5-enoic acid (8-R)

HO COOH

OTBS TIPSO OTIPS A suspension of (4-carboxybutyl)-triphenylphosphonium bromide (1.30 g , 2.94 mmol) in anhydrous THF (100 mL) was cooled to 0 ºC and potassium tert- butoxide (1.0 M in THF, 2.94 mL, 2.94 mmol) was added dropwise. The orange reaction mixture was stirred for 30 min at 0 ºC at which point lactol 7-R (418.8 mg, 0.587 mmol) in anhydrous THF (10mL) was added dropwise. The reaction continued for 1hr and was quenched with saturated NH4Cl and warmed to rt. After separation the organic layer was evaporated, diluted with EtOAc, washed with water and brine, dried over MgSO4, filtered, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 50:50) afforded 364.8 mg (78%) of the desired 1 product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.52 – 5.40 (m, 3H), 5.39 – 5.31 (m, 1H), 4.24 – 4.12 (m, 3H), 3.60 (t, J = 6.53 Hz, 2H), 2.43 – 2.30 (m, 4H), 2.27 – 2.09 (m, 3H), 2.00 (d, J = 13.55 Hz, 1H), 1.80 (dt, J = 13.93 Hz, 4.08 Hz, 1H), 1.71 (quin, J = 7.28 Hz, 2H), 1.61 – 1.44 (m, 5H), 1.36 – 1.23 (m, 4H), 1.11 – 1.01 (m, 42H), 0.89 (s, 9H), 0.05 (s, 6H). HRMS calcd for + C44H88O6NaSi3, [M+Na ] 819.5786, found 819.5784.

(Z)-methyl 7-((1R,2R,3R,5S)-2-((S,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-hydroxy-3- (triisopropylsilyloxy)cyclopentyl)hept-5-enoate (9-S)

HO COOMe

OTBS TIPSO OTIPS To a solution of carboxylic acid 8-S (361.0 mg, 0.45 mmol) in anhydrous diethyl ether (80 mL) and anhydrous MeOH (20 mL) was added tert- butyldimethylsilyl-diazomethane (2.0 M in diethyl ether, 1.13 mL, 2.26 mmol) at rt. The reaction was stirred for 15 min, concentrated, and purified by flash chromatography on silica gel (EtOAc/Hexanes, 40:60). The desired product was 1 isolated as a colorless oil, 342.4 mg (93%). H NMR (400 MHz, CDCl3) δ [ppm] = 5.51 – 5.29 (m, 4H), 4.21 – 4.08 (m, 3H), 3.66 (s, 3H), 3.59 (t, J = 6.51 Hz, 2H), 2.45 – 2.27 (m, 4H), 2.24 – 2.08 (m, 3H), 1.98 (d, J = 13.72 Hz, 1H), 1.81 (dt, J = 13.72 Hz, 4.46 Hz, 1H), 1.70 (quin, J = 7.46 Hz, 2H), 1.61 – 1.42 (m, 5H), 1.36 –

-255-

1.24 (m, 4H), 1.12-1.01 (m, 42H), 0.89 (s, 9H), 0.04 (s, 6H). HRMS calcd for + C45H91O6Si3, [M+H ] 811.6124, found 811.6135.

(Z)-methyl 7-((1R,2R,3R,5S)-2-((R,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-hydroxy-3- (triisopropylsilyloxy)cyclopentyl)hept-5-enoate (9-R)

HO COOMe

OTBS TIPSO OTIPS

To a solution of carboxylic acid 8-R (346.7 mg, 0.43 mmol) in anhydrous diethyl ether (80 mL) and anhydrous MeOH (20 mL) was added tert- butyldimethylsilyl-diazomethane (2.0 M in diethyl ether, 1.08 mL, 2.17 mmol) at rt. The reaction was stirred for 15 min, concentrated, and purified by flash chromatography on silica gel (EtOAc/Hexanes, 40:60). The desired product was 1 isolated as a colorless oil, 321.4 mg (92%). H NMR (400 MHz, CDCl3) δ [ppm] = 5.51 – 5.40 (m, 3H), 5.39 – 5.29 (m, 1H), 4.24 – 4.08 (m, 3H), 3.66 (s, 3H), 3.59 (t, J = 6.51 Hz, 2H), 2.43 – 2.28 (m, 4H), 2.25 – 2.07 (m, 3H), 1.99 (d, J = 13.72 Hz, 1H), 1.80 (dt, J = 13.8 Hz, 4.41 Hz, 1H), 1.69 (quin, J = 7.46 Hz, 2H), 1.60 – 1.42 (m, 5H), 1.35 – 1.23 (m, 4H), 1.12 – 0.99 (m, 42H), 0.89 (s, 9H), 0.04 (s, + 6H). HRMS calcd for C45H91O6Si3, [M+H ] 811.6124, found 811.6134.

(Z)-methyl 7-((1R,2R,3R)-2-((S,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-oxo-3- (triisopropylsilyloxy)cyclopentyl)hept-5-enoate (10-S)

O COOMe

OTBS TIPSO OTIPS A solution of alcohol 9-S (342.4 mg, 0.42 mmol) in anhydrous DCM (30 mL) was cooled to 0 ºC and pyridine (0.65mL, 8.04 mmol) was added followed by Dess- Martin periodinane (358.0 mg, 0.844 mmol). After 10 min, the ice bath was removed and the reaction mixture was stirred and allow to come to rt over 1 hr. It was quenched with NaHCO3:NaS2O3 (1:1), washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 326.7 mg (96%) of the desired product as a 1 colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.61-5.49 (m, 2H), 5.46 – 5.33 (m, 2H), 4.25 – 4.16 (m, 2H), 3.66 (s, 3H), 3.59 (t, J = 6.53 Hz, 2H), 2.69 – 2.55 (m, 2H), 2.46 – 2.33 (m, 2H), 2.30 (t, J = 7.53 Hz, 2H), 2.23 (dd, J = 18.07 Hz, 6.53 Hz, 1H), 2.12 – 2.04 (m, 3H), 1.68 (quin, J = 7.53 Hz, 2H), 1.60 – 1.45 (m, 4H), 1.37 – 1.28 (m, 4H), 1.08 – 1.02 (m, 42H), 0.89 (s, 9H), 0.04 (s, 6H). HRMS + calcd for C45H88O6NaSi3, [M+Na ] 831.5786, found 831.5764.

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(Z)-methyl 7-((1R,2R,3R)-2-((R,E)-8-(tert-butyldimethylsilyloxy)-3- (triisopropylsilyloxy)oct-1-enyl)-5-oxo-3- (triisopropylsilyloxy)cyclopentyl)hept-5-enoate (10-R)

O COOMe

OTBS TIPSO OTIPS

A solution of alcohol 9-R (321.4 mg, 0.40 mmol) in anhydrous DCM (30 mL) was cooled to 0 ºC and pyridine (0.65 mL, 8.0 mmol) was added followed by Dess- Martin periodinane (330.1 mg, 0.80 mmol). After 10 min, the ice bath was removed and the reaction mixture was stirred and allow to come to rt over 1 hr. It was quenched with NaHCO3:NaS2O3 (1:1), washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 313.9 mg (96%) of the desired product as a 1 colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.66 – 5.53 (m, 2H), 5.46 – 5.31 (m, 2H), 4.28 – 4.20 (m, 2H), 3.66 (s, 3H), 3.59 (t, J = 6.53 Hz, 2H), 2.69 – 2.57 (m, 2H), 2.45 – 2.34 (m, 2H), 2.30 (t, J = 7.53 Hz, 2H), 2.23 (dd, J = 17.82 Hz, 6.27 Hz, 1H), 2.11 – 2.01 (m, 3H), 1.68 (quin, J = 7.53 Hz, 2H), 1.60 – 1.44 (m, 4H), 1.37 – 1.25 (m, 4H), 1.10 – 1.09 (m, 42H), 0.89 (s, 9H), 0.04 (s, 6H). HRMS + calcd for C45H88O6NaSi3, [M+Na ] 831.5786, found 831.5790.

(Z)-methyl 7-((1R,2R,3R)-2-((S,E)-8-hydroxy-3-(triisopropylsilyloxy)oct-1- enyl)-5-oxo-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoate (11-S)

O COOMe

OH TIPSO OTIPS To a solution of TBS-protected 10-S (326.7 mg, 0.40 mmol) in THF (2 mL) was added H2O (2mL) and AcOH (6mL). The reaction mixture was stirred at rt overnight. It was quenched with NaHCO3, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 50:50) afforded 262.2 mg (93%) of the desired product as a 1 colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.61 – 5.51 (m, 2H), 5.47 – 5.33 (m, 2H), 4.27 – 4.16 (m, 2H), 3.69 – 3.59 (m, 5H), 2.70 – 2.55 (m, 2H), 2.46 – 2.33 (m, 2H), 2.30 (t, J = 7.78 Hz, 2H), 2.23 (dd, J = 17.82 Hz, 6.27 Hz, 1H), 2.12 – 2.03 (m, 3H), 1.68 (quin, J = 7.53 Hz, 2H), 1.61 – 1.48 (m, 4H), 1.39 – 1.29 (m, + 4H), 1.09 – 1.00 (m, 42H). HRMS calcd for C39H74O6NaSi2, [M+Na ] 717.4922, found 717.4946.

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(Z)-methyl 7-((1R,2R,3R)-2-((R,E)-8-hydroxy-3-(triisopropylsilyloxy)oct-1- enyl)-5-oxo-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoate (11-R)

O COOMe

OH TIPSO OTIPS

To a solution of TBS-protected 10-R (313.9, 0.39 mmol) in THF (2 mL) was added H2O (2mL) and AcOH (6mL). The reaction mixture was stirred at rt overnight. It was quenched with NaHCO3, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 50:50) afforded 245.3 mg (91%) of the desired product as a colorless oil. 1H NMR (400 MHz, CDCl3) δ [ppm] = 5.66 – 5.53 (m, 2H), 5.46 – 5.32 (m, 2H), 4.28 – 4.21 (m, 2H), 3.69 – 3.60 (m, 5H), 2.69 – 2.57 (m, 2H), 2.45 – 2.34 (m, 2H), 2.30 (t, J = 7.53 Hz, 2H), 2.24 (dd, J = 18.07 Hz, 6.53 Hz, 1H), 2.11 – 2.01 (m, 3H), 1.68 (quin, J = 7.53 Hz, 2H), 1.62 – 1.48 (m, 4H), 1.39 – 1.29 (m, 4H), 1.11 – 1.00 (m, + 42H). HRMS calcd for C39H74O6NaSi2, [M+Na ] 717.4922, found 717.4904.

(Z)-methyl 7-((1R,2R,3R)-2-((S,E)-8-azido-3-(triisopropylsilyloxy)oct-1- enyl)-5-oxo-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoate (12-S)

O COOMe

N3 TIPSO OTIPS To a solution of alcohol 11-S (50 mg, 0.072 mmol) and triphenylphosphine (27.7 mg, 0.144 mmol) in anhydrous toluene (2.0 mL) was added Zn(N3)2-2Py (26.4 mg, 0.086 mmol). Diisopropyl azodicarboxylate (28 µL, 0.144 mmol) was then added and the reaction mixture was allowed to continue stirring for 7 hrs. The mixture was concentrated, and purification by flash chromatography on silica gel (EtOAc/Hexanes, 90:10) afforded 47.3 mg (91%) of the desired product as a colorless oil. 1H NMR (400 MHz, CDCl3) δ [ppm] = 5.61 – 5.50 (m, 2H), 5.47 – 5.32 (m, 2H), 4.27 – 4.16 (m, 2H), 3.66 (s, 3H), 3.26 (t, J = 7.03 Hz, 2H), 2.69 – 2.55 (m, 2H), 2.47 – 2.33 (m, 2H), 2.30 (t, J = 7.54 Hz, 2H), 2.23 (dd, J = 18.00 Hz, 6.69 Hz, 1H), 2.12 – 2.04 (m, 3H), 1.68 (quin, J = 7.54 Hz, 2H), 1.63 – 1.48 (m, 4H), 1.42 – 1.31 (m, + 4H), 1.13 – 1.00 (m, 42H). HRMS calcd for C39H73N3O5NaSi2, [M+Na ] 717.4986, found 742.4991.

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(Z)-methyl 7-((1R,2R,3R)-2-((R,E)-8-azido-3-(triisopropylsilyloxy)oct-1- enyl)-5-oxo-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoate (12-R)

O COOMe

N3 TIPSO OTIPS

To a solution of alcohol 11-R (62 mg, 0.089 mmol) and triphenylphosphine (46.5 mg, 0.178 mmol) in anhydrous toluene (2.0 mL) was added Zn(N3)2-2Py (32.5 mg, 0.106 mmol). Diisopropyl azodicarboxylate (35 µL, 0.178 mmol) was then added and the reaction mixture was allowed to continue stirring for 7 hrs. The mixture was concentrated, and purification by flash chromatography on silica gel (EtOAc/Hexanes, 90:10) afforded 56.2 (91%) of the desired product as a colorless oil. 1H NMR (400 MHz, CDCl3) δ [ppm] = 5.67 – 5.53 (m, 2H), 5.46 – 5.31 (m, 2H), 4.29 – 4.21 (m, 2H), 3.66 (s, 3H), 3.25 (t, J = 6.86 Hz, 2H), 2.69 – 2.57 (m, 2H), 2.46 – 2.33 (m, 2H), 2.30 (t, J = 7.72 Hz, 2H), 2.23 (dd, J = 18.00 Hz, 6.34 Hz, 1H), 2.11 – 2.02 (m, 3H), 1.68 (quin, J = 7.54 Hz, 2H), 1.62 – 1.48 (m, 4H), 1.39 – 1.29 (m, 4H), + 1.10 – 1.01 (m, 42H). HRMS calcd for C39H73N3O5NaSi2, [M+Na ] 717.4986, found 742.4997.

(Z)-methyl 7-((1R,2R,3R)-2-((S,E)-8-azido-3-(triisopropylsilyloxy)oct-1- enyl)-5-hydroxy-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoate (13-S) (+)/(-)

HO COOMe

N3 TIPSO OTIPS To a solution of azide 12-S (64.4 mg, 0.089 mmol) in THF (0.5mL) and MeOH (3.0mL) at 0 ºC was added cerium (III) chloride heptahydrate (33.2 mg, 0.089 mmol) and sodium borohydride (3.4 mg, 0.089 mmol). After 1hr stirring, the reaction was concentrated, diluted with EtOAc, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 53.9 mg (83%) of the desired product as a 1 colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.51 – 5.30 (m, 4H), 4.22 – 4.08 (m, 3H), 3.66 (s, 3H), 3.25 (t, J = 7.03 Hz, 2H), 2.45 – 2.28 (m, 4H), 2.25 – 2.08 (m, 3H), 2.02 – 1.94 (m, 1H), 1.86 – 1.77 (m, 1H), 1.70 (quin, J = 7.46 Hz, 2H), 1.63 – 1.47 (m, 5H), 1.40 – 1.28 (m, 4H), 1.11 – 1.00 (m, 42H). HRMS calcd for + C39H76N3O5Si2, [M+H ] 722.5324, found 722.5320.

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(Z)-methyl 7-((1R,2R,3R)-2-((R,E)-8-azido-3-(triisopropylsilyloxy)oct-1- enyl)-5-hydroxy-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoate (13-R) (+)/(-)

HO COOMe

N3 TIPSO OTIPS To a solution of azide 12-R (70.7 mg, 0.098 mmol) in THF (0.5mL) and MeOH (3.0mL) at 0 ºC was added cerium (III) chloride heptahydrate (36.5 mg, 0.098 mmol) and sodium borohydride (3.7 mg, 0.098 mmol). After 1hr stirring, the reaction was concentrated, diluted with EtOAc, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 63.2 mg (89%) of the desired product as a 1 colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.51 – 5.30 (m, 4H), 4.23 – 4.08 (m, 3H), 3.66 (s, 3H), 3.25 (t, J = 6.86 Hz, 2H), 2.44 – 2.28 (m, 4H), 2.25 – 2.08 (m, 3H), 2.04 – 1.95 (m, 1H), 1.86 – 1.77 (m, 1H), 1.70 (quin, J = 7.54 Hz, 2H), 1.62 – 1.47 (m, 5H), 1.40 – 1.28 (m, 4H), 1.11 – 1.00 (m, 42H). HRMS calcd for + C39H76N3O5Si2, [M+H ] 722.5324, found 722.5313.

(Z)-7-((1R,2R,3R)-2-((S,E)-8-azido-3-(triisopropylsilyloxy)oct-1-enyl)-5- hydroxy-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoic acid (14-S) (+)/(- )

HO COOH

N3 TIPSO OTIPS Aqueous NaOH (1M, 2.5 mL) was added to a solution of ester 13-S (893.6 mg, 1.24 mmol) in THF (10mL). The reaction was stirred for 24 hrs and neutralized to pH5 using citric acid. The mixture was concentrated down to just water, extracted with DCM, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 90:10) afforded 1 741.5mg (85%) of the desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.52 – 5.30 (m, 4H), 4.22 – 4.11 (m, 3H), 3.25 (t, J = 6.86 Hz, 2H), 2.46 – 2.31 (m, 4H), 2.25 – 2.10 (m, 3H), 2.04 – 1.95 (m, 1H), 1.85 – 1.77 (m, 1H), 1.71 (quin, J = 7.37 Hz, 2H), 1.63 – 1.47 (m, 5H), 1.40 – 1.28 (m, 4H), 1.11 – 1.00 (m, + 42H). HRMS calcd for C38H73N3O5NaSi2, [M+Na ] 730.4986, found 730.4995.

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(Z)-7-((1R,2R,3R)-2-((R,E)-8-azido-3-(triisopropylsilyloxy)oct-1-enyl)-5- hydroxy-3-(triisopropylsilyloxy)cyclopentyl)hept-5-enoic acid (14-R) (+)/(- )

HO COOH

N3 TIPSO OTIPS

Aqueous NaOH (1M, 2.5 mL) was added to a solution of ester 13-R (672.7, 0.93 mmol) in THF (10mL). The reaction was stirred for 24 hrs and neutralized to pH5 using citric acid. The mixture was concentrated down to just water, extracted with DCM, washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 90:10) afforded 1 659.5 mg (84%) of the desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.51 – 5.31 (m, 4H), 4.22 – 4.12 (m, 3H), 3.25 (t, J = 6.86 Hz, 2H), 2.45 – 2.31 (m, 4H), 2.25 – 2.10 (m, 3H), 2.05 – 1.96 (m, 1H), 1.85 – 1.77 (m, 1H), 1.71 (quin, J = 7.29 Hz, 2H), 1.63 – 1.47 (m, 5H), 1.40 – 1.28 (m, 4H), 1.11 – 1.00 + (m, 42H). HRMS calcd for C38H73N3O5NaSi2, [M+Na ] 730.4986, found 730.4997.

General procedure for amide syntheses

To a solution of the carboxylic acid 14 (1.0 equiv) in anhydrous DCM was added carbonyldiimidazole (3.0 equiv) at rt. The reaction was allowed to stir for 30 min, and the desired amine H2N-R1 (4.0 equiv) was added. The reaction was allowed to progress for 3 hrs at which point the mixture was concentrated and purified by flash chromatography on silica gel.

(Z)-7-((1R,2R,3R)-2-((S,E)-8-azido-3-(triisopropylsilyloxy)oct-1-enyl)-5- hydroxy-3-(triisopropylsilyloxy)cyclopentyl)-N-(2-(tert- butyldimethylsilyloxy)ethyl)hept-5-enamide (15-S) (+)/(-)

HO H N OTBS O N3 TIPSO OTIPS

To a solution of the carboxylic acid 14-S (687.0 mg, 0.97 mmol) in anhydrous DCM (35 mL) was added carbonyldiimidazole (472 mg, 2.91 mmol) at rt. The reaction was allowed to stir for 30 min, and a solution of TBS protected ethanolamine (681 mg, 3.88 mmol) in anhydrous DCM (3.0 mL) was added and allowed to react for another 3 hrs. The mixture was concentrated, and purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 738.6mg (88%) 1 of the desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.91 (br. s., 1H), 5.49 – 5.31 (m, 4H), 4.23 – 4.08 (m, 3H), 3.67 (t, J = 5.31, 2H), 3.36

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(q, J = 5.03 Hz, 2H), 3.25 (t, J = 7.03 Hz, 2H), 2.48 – 2.32 (m, 2H), 2.23 – 2.05 (m, 5H), 2.02 – 1.94 (m, 1H), 1.87 – 1.78 (m, 1H), 1.71 (quin, J = 7.23, 2H), 1.63 – 1.45 (m, 5H), 1.41 – 1.26 (m, 4H), 1.12 – 1.00 (m, 42H), 0.90 (s, 9H), 0.06 (s, + 6H). HRMS calcd for C46H93N4O5Si2, [M+H ] 865.6454, found 865.6450.

(Z)-7-((1R,2R,3R)-2-((R,E)-8-azido-3-(triisopropylsilyloxy)oct-1-enyl)-5- hydroxy-3-(triisopropylsilyloxy)cyclopentyl)-N-(2-(tert- butyldimethylsilyloxy)ethyl)hept-5-enamide (15-R) (+)/(-)

HO H N OTBS O N3 TIPSO OTIPS To a solution of the carboxylic acid 14-R (659.5 mg, 0.93 mmol) in anhydrous DCM (35 mL) was added carbonyldiimidazole (453 mg, 2.79 mmol) at rt. The reaction was allowed to stir for 30 min, and a solution of TBS protected ethanolamine (652.3 mg, 3.72 mmol) in anhydrous DCM (3.0 mL) was added and allowed to react for another 3 hrs. The mixture was concentrated, and purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 741.4 mg 1 (92%) of the desired product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.91 (br. s., 1H), 5.49 – 5.30 (m, 4H), 4.23 – 4.07 (m, 3H), 3.67 (t, J = 5.31, 2H), 3.36 (q, J = 4.98 Hz, 2H), 3.25 (t, J = 6.86 Hz, 2H), 2.47 – 2.32 (m, 2H), 2.23 – 2.04 (m, 5H), 2.02 – 1.94 (m, 1H), 1.86 – 1.78 (m, 1H), 1.71 (quin, J = 7.20, 2H), 1.63 – 1.45 (m, 5H), 1.41 – 1.26 (m, 4H), 1.12 – 1.00 (m, 42H), 0.90 (s, 9H), 0.06 + (s, 6H). HRMS calcd for C46H93N4O5Si2, [M+H ] 865.6454, found 865.6456.

(Z)-7-((1R,2R,3R)-2-((S,E)-8-azido-3-(triisopropylsilyloxy)oct-1-enyl)-5- oxo-3-(triisopropylsilyloxy)cyclopentyl)-N-(2-(tert- butyldimethylsilyloxy)ethyl)hept-5-enamide (16-S)

O H N OTBS O N3 TIPSO OTIPS

A solution of alcohol 15-S (738.6 mg, 0.98 mmol) in anhydrous DCM (50mL) was cooled to 0 ºC. Pyridine (1.6 mL, 19.6 mmol) was added followed by dess- martin periodinane (834 mg, 1.96 mmol) and the mixture was stirred for 1 hr slowly coming to rt. Quenching with NaHCO3:NaS2O3 (1:1) was followed by extraction with DCM. The mixture was dried over MgSO4, and purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 719.4 mg (98%) of the desired 1 product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.88 (br. s., 1H), 5.67 – 5.50 (m, 2H), 5.47 – 5.31 (m, 2H), 4.29 – 4.16 (m, 2H), 3.67 (t, J = 5.31 Hz, 2H), 3.37 (q, J = 5.49 Hz, 2H), 3.25 (t, J = 6.86 Hz, 2H), 2.70 – 2.56 (m, 2H), 2.48 – 2.29 (m, 2H), 2.27 – 2.14 (m, 3H), 2.12 – 2.03 (m, 3H), 1.75 – 1.65 (m, 2H), 1.61 – 1.48 (m, 4H), 1.41 – 1.30 (m, 4H), 1.11 – 1.00 (m, 42H), 0.90 (s, 9H), + 0.06 (s, 6H). HRMS calcd for C46H91N4O5Si2, [M+H ] 863.6297, found 863.6288.

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(Z)-7-((1R,2R,3R)-2-((R,E)-8-azido-3-(triisopropylsilyloxy)oct-1-enyl)-5- oxo-3-(triisopropylsilyloxy)cyclopentyl)-N-(2-(tert- butyldimethylsilyloxy)ethyl)hept-5-enamide (16-R)

O H N OTBS O N3 TIPSO OTIPS

A solution of alcohol 15-R (741.4 mg, 0.99 mmol) in anhydrous DCM (50mL) was cooled to 0 ºC. Pyridine (1.59 mL, 19.7 mmol) was added followed by dess- martin periodinane (836 mg, 1.97 mmol) and the mixture was stirred for 1 hr slowly coming to rt. Quenching with NaHCO3:NaS2O3 (1:1) was followed by extraction with DCM. The mixture was dried over MgSO4, and purification by flash chromatography on silica gel (EtOAc/Hexanes, 30:70) afforded 718.8 mg (97%) of the desired 1 product as a colorless oil. H NMR (400 MHz, CDCl3) δ [ppm] = 5.88 (br. s., 1H), 5.67 – 5.52 (m, 2H), 5.47 – 5.31 (m, 2H), 4.29 – 4.16 (m, 2H), 3.67 (t, J = 5.14 Hz, 2H), 3.37 (q, J = 5.49 Hz, 2H), 3.25 (t, J = 6.86 Hz, 2H), 2.70 – 2.56 (m, 2H), 2.48 – 2.29 (m, 2H), 2.27 – 2.14 (m, 3H), 2.12 – 2.02 (m, 3H), 1.76 – 1.64 (m, 2H), 1.62 – 1.50 (m, 4H), 1.39 – 1.29 (m, 4H), 1.10 – 1.01 (m, 42H), 0.90 (s, 9H), + 0.07 (s, 6H). HRMS calcd for C46H91N4O5Si2, [M+H ] 863.6297, found 863.6307.

(Z)-7-((1R,2R,3R)-2-((S,E)-8-azido-3-hydroxyoct-1-enyl)-3-hydroxy-5- oxocyclopentyl)-N-(2-hydroxyethyl)hept-5-enamide (AM7638)

O H N OH O N3 H O OH A Teflon vial was charged with silyl protected 16-S (21 mg, 0.024 mmol) in THF (0.2 mL) and CH3CN (0.3 mL). The vial was cooled to 0 ºC and pyridine (20 µL) was added followed by 48% HF (50 µL). The reaction was stirred overnight slowly coming to rt. After 24 hrs the mixture was poured into chloroform (4 mL), quenched with saturated NaHCO3, and washed with water and brine. The organic layers were combined, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (Acetone/Hexanes, 90:10) afforded 9.0 mg (85%) of the desired product 1 as a colorless oil. H NMR (500 MHz, CDCl3) δ [ppm] = 6.34 (br. s., 1H), 5.71 (dd, J = 15.14 Hz, 6.84 Hz, 1H), 5.58 (dd, J = 15.63 Hz, 8.30 Hz, 1H), 5.47 – 5.38 (m, 1H), 5.38 – 5.29 (m, 1H), 4.15 – 4.06 (m, 2H), 3.69 (br. s., 2H), 3.40 (q, J = 5.37 Hz, 2H), 3.28 (t, J = 6.84 Hz, 2H), 2.86 (br. s., 1H), 2.75 (dd, J = 18.31 Hz, 7.08 Hz, 1H), 2.48 – 2.39 (m, 2H), 2.30 (dt, J = 14.16 Hz, 5.86 Hz, 1H), 2.25 – 2.17 (m, 3H), 2.14 (dt, J = 11.11 Hz, 5.43 Hz, 1H), 2.06 (q, J = 7.32 Hz, 2H), 1.82 (br. s., 13 1H), 1.75 – 1.50 (m, 6H), 1.49 – 1.34 (m, 4H). C NMR (100 MHz, CDCl3) δ [ppm] = 214.92, 174.52, 137.43, 131.23, 129.94, 126.45, 72.30, 72.13, 62.09, 54.64, 52.80, 51.36, 46.52, 42.27, 37.12, 35.83, 28.77, 26.70, 26.61, 25.46, 25.14, 24.83. + HRMS calcd for C22H37N4O5, [M+H ] 437.2764, found 437.2751.

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(Z)-7-((1R,2R,3R)-2-((R,E)-8-azido-3-hydroxyoct-1-enyl)-3-hydroxy-5- oxocyclopentyl)-N-(2-hydroxyethyl)hept-5-enamide (AM7637)

O H N OH O N3 H O OH A Teflon vial was charged with silyl protected 16-R (27 mg, 0.031 mmol) in THF (0.2 mL) and CH3CN (0.3 mL). The vial was cooled to 0 ºC and pyridine (20 µL) was added followed by 48% HF (50 µL). The reaction was stirred overnight slowly coming to rt. After 24 hrs the mixture was poured into chloroform (4 mL), quenched with saturated NaHCO3, and washed with water and brine. The organic layers were combined, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (Acetone/Hexanes, 90:10) afforded 12.0 mg (88%) of the desired 1 product as a colorless oil. H NMR (500 MHz, CDCl3) δ [ppm] = 6.32 (t, J = 5.13 Hz, 1H), 5.66 (dd, J = 15.63 Hz, 6.84 Hz, 1H), 5.59 (dd, J = 15.63 Hz, 7.81 Hz, 1H), 5.45 – 5.38 (m, 1H), 5.36 – 5.28 (m, 1H), 4.14 – 4.04 (m, 2H), 3.70 (br. s., 2H), 3.40 (q, J = 5.37 Hz, 2H), 3.28 (t, J = 6.84 Hz, 2H), 2.74 (dd, J = 18.56 Hz, 7.32 Hz, 1H), 2.48 – 2.36 (m, 2H), 2.30 (dt, J = 14.65 Hz, 6.84 Hz, 1H), 2.25 – 2.17 (m, 3H), 2.13 (dt, J = 11.48 Hz, 5.49 Hz, 1H), 2.06 (q, J = 6.90 Hz, 2H), 1.81 (br. s., 1H), 1.74 – 1.65 (m, 2H), 1.65 – 1.55 (m, 3H) 1.55 – 1.47 (m, 1H), 1.47 – 1.33 (m, 13 4H). C NMR (100 MHz, CDCl3) δ [ppm] = 215.20, 174.44, 136.76, 131.14, 130.95, 126.53, 72.59, 72.17, 61.86, 54.30, 53.13, 51.38, 46.39, 42.28, 36.90, 35.77, + 28.83, 26.64, 25.43, 25.14. HRMS calcd for C22H37N4O5, [M+H ] 437.2764, found 437.2750.

(Z)-7-((1R,2R,3R)-2-((S,E)-8-isothiocyanato-3-hydroxyoct-1-enyl)-3- hydroxy-5-oxocyclopentyl)-N-(2-hydroxyethyl)hept-5-enamide (AM7645)

O H N OH O NCS H O OH To a solution of azide AM7638 (12 mg, 0.027 mmol) in anhydrous THF (2mL) was added triphenylphosphine (11 mg, 0.041mmol) at rt. Carbon disulfide (17 µL, 0.280 mmol) was added, the reaction was stirred for 48 hrs, and then concentrated. Purification of the residue by flash chromatography on silica gel (Acetone/Hexanes, 80:20) afforded 11.9 mg (96%) of the desired product as a colorless oil. 1H NMR (500 MHz, CDCl3) δ [ppm] = 6.23 (br. s., 1H), 5.71 (dd, J = 15.14 Hz, 6.84 Hz, 1H), 5.58 (dd, J = 15.14 Hz, 8.30 Hz, 1H), 5.48 – 5.39 (m, 1H), 5.39 – 5.29 (m, 1H), 4.17 – 4.07 (m, 2H), 3.70 (t, J = 4.88, 2H), 3.53 (t, J = 6.59 Hz, 2H), 3.41 (q, J = 5.37 Hz, 2H), 2.76 (dd, J = 18.31 Hz, 7.08 Hz, 1H), 2.48 – 2.40 (m, 2H), 2.30 (dt, J = 14.65 Hz, 6.84 Hz, 1H), 2.25 – 2.17 (m, 3H), 2.14 (dt, J = 11.35 Hz, 5.31 Hz, 1H), 2.06 (q, J = 7.32 Hz, 2H), 1.76 – 1.57 (m, 7H), 1.51 – 1.42 (m, 3H). HRMS + calcd for C23H37N2O5S, [M+H ] 453.2423, found 453.2427.

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(Z)-7-((1R,2R,3R)-2-((R,E)-8-isothiocyanato-3-hydroxyoct-1-enyl)-3- hydroxy-5-oxocyclopentyl)-N-(2-hydroxyethyl)hept-5-enamide (AM7646)

O H N OH O NCS H O OH To a solution of azide AM7637 (16 mg, 0.037 mmol) in anhydrous THF (2mL) was added triphenylphosphine (14.7 mg, 0.056 mmol) at rt. Carbon disulfide (22 µL, 0.370 mmol) was added, the reaction was stirred for 48 hrs, and then concentrated. Purification of the residue by flash chromatography on silica gel (Acetone/Hexanes, 80:20) afforded 15.6 mg (94%) of the desired product as a colorless oil. 1H NMR (500 MHz, CDCl3) δ [ppm] = 6.31 (br. s., 1H), 5.67 (dd, J = 15.14 Hz, 6.35 Hz, 1H), 5.61 (dd, J = 15.63 Hz, 7.81 Hz, 1H), 5.45 – 5.38 (m, 1H), 5.36 – 5.29 (m, 1H), 4.14 – 4.05 (m, 2H), 3.70 (t, J = 4.64, 2H), 3.53 (t, J = 6.35 Hz, 2H), 3.41 (q, J = 5.37 Hz, 2H), 2.74 (dd, J = 18.56 Hz, 7.32 Hz, 1H), 2.48 – 2.37 (m, 2H), 2.30 (dt, J = 13.92 Hz, 6.71 Hz, 1H), 2.25 – 2.17 (m, 3H), 2.15 (dt, J = 11.48 Hz, 5.49 Hz, 1H), 2.07 (q, J = 7.02 Hz, 2H), 1.78 (br. s., 1H), 1.75 – 1.65 (m, 4H), 1.63 – 1.56 + (m, 1H), 1.56 – 1.34 (m, 5H). HRMS calcd for C23H37N2O5S, [M+H ] 453.2423, found 453.2428.

(Z)-7-((1R,2R,3R)-3-hydroxy-2-((S,E)-3-hydroxyoct-1-enyl)-5- oxocyclopentyl)-N-((S)-1-hydroxypropan-2-yl)hept-5-enamide (AM7648)

O H N OH O

HO OH 1 H NMR (500 MHz, CDCl3) δ [ppm] = 5.93 (br. s., 1H), 5.67 (dd, J=15.14, 6.35 Hz, 1H), 5.60 (dd, J=15.14, 7.81 Hz, 1H), 5.29 - 5.37 (m, 1H), 4.02 - 4.15 (m, 3H), 3.65 (d, J=9.77 Hz, 1H), 3.48 (dd, J=9.77, 6.84 Hz, 1H), 2.74 (dd, J=18.31, 7.08 Hz, 1H), 2.37 - 2.49 (m, 2H), 2.30 (dt, J=13.92, 6.71 Hz, 1H), 2.10 - 2.26 (m, 4H), 2.01 - 2.10 (m, 2H), 1.68 - 1.79 (m, 4H), 1.60 - 1.68 (m, 1H), 1.26 - 1.37 (m, 6H), + 1.16 (d, J=6.84 Hz, 3H), 0.89 (t, J=6.59 Hz, 3H). HRMS calcd for C23H40NO5, [M+H ] 410.2906, found 410.2897.

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(Z)-7-((1R,2R,3R)-3-hydroxy-2-((S,E)-3-hydroxyoct-1-enyl)-5- oxocyclopentyl)-N-((R)-1-hydroxypropan-2-yl)hept-5-enamide (AM7649)

O H N OH O

HO OH 1 H NMR (500 MHz, CDCl3) δ [ppm] = 5.88 (d, J=5.37 Hz, 1H), 5.67 (dd, J=15.63, 6.84 Hz, 1H), 5.59 (dd, J=15.63, 8.30 Hz, 1H), 5.37 - 5.47 (m, 1H), 5.28 - 5.37 (m, 1H), 4.01 - 4.15 (m, 3H), 3.66 (d, J=10.25 Hz, 1H), 3.48 (dd, J=10.25, 6.35 Hz, 1H), 2.74 (dd, J=18.31, 7.08 Hz, 1H), 2.36 - 2.49 (m, 2H), 2.27 - 2.36 (m, 1H), 2.16 - 2.26 (m, 3H), 1.99 - 2.17 (m, 3H), 1.63 - 1.75 (m, 6H), 1.27 - 1.37 (m, 5H), + 1.16 (d, J=6.84 Hz, 3H), 0.89 (t, J=6.59 Hz, 3H). HRMS calcd for C23H40NO5, [M+H ] 410.2906, found 410.2908.

(Z)-N-cyclopropyl-7-((1R,2R,3R)-3-hydroxy-2-((S,E)-3-hydroxyoct-1-enyl)- 5-oxocyclopentyl)hept-5-enamide (AM7650)

O H N

O

HO OH 1 H NMR (500 MHz, CDCl3) δ [ppm] = 5.85 (br. s., 1H), 5.68 (dd, J=15.14, 6.35 Hz, 1H), 5.59 (dd, J=15.63, 8.30 Hz, 1H), 5.36 - 5.46 (m, 1H), 5.28 - 5.36 (m, 1H), 4.04 - 4.14 (m, 2H), 3.49 (br. s., 1H), 3.05 (br. s., 1H), 2.66 - 2.78 (m, 2H), 2.35 - 2.49 (m, 2H), 2.17 - 2.33 (m, 2H), 2.08 - 2.16 (m, 3H), 1.96 - 2.09 (m, 2H), 1.60 - 1.73 (m, 2H), 1.54 - 1.60 (m, 1H), 1.44 - 1.54 (m, 1H), 1.26 - 1.35 (m, 5H), 0.85 - 0.94 (m, 3H), 0.72 - 0.79 (m, 2H), 0.43 - 0.51 (m, 2H). HRMS calcd for C23H38NO4, [M+H+] 392.2801, found 392.2797.

(Z)-7-((1R,2R,3R)-2-((S,E)-8-azido-3-hydroxyoct-1-enyl)-3-hydroxy-5- oxocyclopentyl)-N-((R)-1-hydroxypropan-2-yl)hept-5-enamide (AM7651)

O H N OH O N3 HO OH 1 H NMR (500 MHz, CDCl3) δ [ppm] = 5.82 (br. s., 1H), 5.57 - 5.73 (m, 2H), 5.28 - 5.48 (m, 2H), 4.02 - 4.19 (m, 3H), 3.66 (d, J=7.81 Hz, 1H), 3.48 (dd, J=10.99, 6.59 Hz, 1H), 3.28 (t, J=6.84 Hz, 2H), 2.74 (dd, J=18.31, 7.08 Hz, 1H), 2.37 - 2.52 (m, 2H), 2.26 - 2.37 (m, 1H), 1.97 - 2.28 (m, 8H), 1.49 - 1.74 (m, 9H), 1.32 - 1.48 (m, + 2H), 1.17 (m, J=6.84 Hz, 3H). HRMS calcd for C23H39N4O5, [M+H ] 451.2920, found 451.2922.

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(Z)-7-((1R,2R,3R)-2-((S,E)-8-azido-3-hydroxyoct-1-enyl)-3-hydroxy-5- oxocyclopentyl)-N-cyclopropylhept-5-enamide (AM7652)

O H N

O N3 HO OH 1 H NMR (500 MHz, CDCl3) δ [ppm] = 5.59 - 5.79 (m, 3H), 5.28 - 5.50 (m, 2H), 4.07 - 4.19 (m, 2H), 3.28 (t, J=6.84 Hz, 2H), 2.65 - 2.84 (m, 3H), 2.36 - 2.52 (m, 2H), 2.30 (dd, J=14.16, 6.35 Hz, 1H), 2.23 (dd, J=18.56, 8.79 Hz, 1H), 2.09 - 2.17 (m, 3H), 1.97 - 2.09 (m, 2H), 1.52 - 1.68 (m, 7H), 1.35 - 1.48 (m, 4H), 0.73 - 0.80 (m, + 2H), 0.44 - 0.50 (m, 2H). HRMS calcd for C23H37N4O4, [M+H ] 433.2815, found 433.2811.

(Z)-1,3-dihydroxypropan-2-yl 7-((1R,2R,3R)-3-hydroxy-2-((S,E)-3- hydroxyoct-1-enyl)-5-oxocyclopentyl)hept-5-enoate (AM7655)

OH O O OH O

HO OH 1 H NMR (500 MHz, CDCl3) δ [ppm] = 5.71 (dd, J=15.63, 6.35 Hz, 1H), 5.61 (dd, J=15.14, 8.30 Hz, 1H), 5.38 - 5.46 (m, 1H), 5.30 - 5.38 (m, 1H), 4.88 - 4.97 (m, 1H), 4.04 - 4.19 (m, 2H), 3.81 - 3.87 (m, 4H), 2.76 (dd, J=18.31, 7.08 Hz, 1H), 2.27 - 2.50 (m, 6H), 2.05 - 2.26 (m, 5H), 1.65 - 1.75 (m, 2H), 1.31 (br. s., 6H), + 0.85 - 0.94 (m, 3H). HRMS calcd for C23H39O7, [M+H ] 427.2696, found 427.2699.

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Appendix 2: Biological Experimental Procedures and Supplemental Data for Assays Presented in Chapter 3

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Experimental Procedure and Data Analysis for Primary Screening in the Psychoactive Drug Screening Program at the University of North Carolina – Chapel Hill:

Test and reference compounds (see Table 1) are diluted to 5X final assay concentration (50 μM for a final assay concentration of 10 μM) in the appropriate radioligand binding buffer (see Table 1). Then, 50-μl aliquots of buffer (negative control), test compound, and reference compound are added in quadruplicate to the wells of a 96-well plate, each of which contains 50 μl of 5X radioligand (see Table 1 for final assay concentration for each radioligand) and 100 μl of buffer (see Table 1). Finally, receptor-containing, crude membrane fractions (prepared as detailed below) are resuspended in an appropriate volume of buffer (see Table 1) and dispensed (50 μl per well) into 96-well plate. Radioligand binding is allowed to equilibrate (typically for 1.5 hours at room temperature, see Table 1), and then bound radioactivity is isolated by filtration onto 0.3% polyethyleneimine-treated, 96-well filter mats using a 96-well Filtermate harverster. The filter mats are dried, then scintillant is melted onto the filters and the radioactivity retained on the filters is counted in a Microbeta scintillation counter.

Table 1 Assay conditions for primary radioligand binding assays. Receptor Radioligand Reference Assay Buffer

(Assay Conc.)

5-HT1A [3H]8-OH-DPAT Methysergide Standard Binding (0.5 nM) Buffer

5-HT1B [3H]GR127543 Ergotamine Standard Binding (0.3 nM) Buffer

5-HT1D [3H]GR127543 Ergotamine Standard Binding (0.3 nM) Buffer

5-HT1E [3H]5-HT (3 nM) 5-HT Standard Binding Buffer

5-HT2A [3H]Ketanserin Chlorpromazine Standard Binding (0.5 nM) Buffer

5-HT2B [3H]LSD (1 nM) Methysergide Standard Binding Buffer

5-HT2C [3H]Mesulergine Chlorpromazine Standard Binding (0.5 nM) Buffer

5-HT3 [3H]LY278584 LY278584 Standard Binding (0.3 nM) Buffer

5-HT5a [3H]LSD (1 nM) Ergotamine Standard Binding Buffer

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5-HT6 [3H]LSD (1 nM) Chlorpromazine Standard Binding Buffer

5-HT7 [3H]LSD (1 nM) Chlorpromazine Standard Binding Buffer

D1 [3H]SCH233930 SKF38393 Dopamine Binding (0.2 nM) Buffer

D2 [3H]N- Haloperidol Dopamine Binding methylspiperone Buffer (0.2 nM)

D3 [3H]N- Chlorpromazine Dopamine Binding methylspiperone Buffer (0.2 nM)

D4 [3H]N- Chlorpromazine Dopamine Binding methylspiperone Buffer (0.3 nM)

D5 [3H]SCH233930 SKF38393 Dopamine Binding (0.2 nM) Buffer

Delta OR [3H]DADLE (0.3 Naltrindole Standard Binding nM) Buffer

Kappa OR [3H]U69593 (0.3 Salvinorin A Standard Binding nM) Buffer

Mu OR [3H]DAMGO (0.3 DAMGO Standard Binding nM) Buffer

H1 [3H]Pyrilamine Chlorpheniramine Histamine Binding (0.9 nM) Buffer

H2 [3H]Tiotidine (3 Cimetidine Histamine Binding nM) Buffer

H3 [3H]alpha- Histamine Histamine Binding methylhistamine Buffer (0.4 nM)

H4 [3H]Histamine (5 Clozapine Histamine Binding nM) Buffer

SERT [3H]Citalopram Amitriptyline Transporter (0.5 nM) Binding Buffer

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NET [3H]Nisoxetine Desipramine Transporter (0.5 nM) Binding Buffer

DAT [3H]WIN35428 GBR12909 Transporter (0.5 nM) Binding Buffer

GABAA [3H]Muscimol (1 GABA 50 mM Tris nM) Acetate, pH 7.4

BZP [3H]Flunitrazepam Diazepam 50 mM Tris HCl, (0.5 nM) 2.5 mM CaCl2, pH 7.4

Alpha1A [3H]Prazosin (0.7 Urapidil Alpha1 Binding nM) Buffer

Alpha1B [3H]Prazosin (0.7 Corynanthine Alpha1 Binding nM) Buffer

Alpha2A [3H]Clonidine (1 Oxymetazoline Alpha2 Binding nM) Buffer

Alpha2B [3H]Clonidine (1 Prazosin Alpha2 Binding nM) Buffer

Alpha2C [3H]Clonidine (1 Prazosin Alpha2 Binding nM) Buffer

Beta1 [125I]Iodopindolol Atenolol Beta Binding (0.1 nM) Buffer

Beta2 [125I]Iodopindolol ICI118551 Beta Binding (0.1 nM) Buffer

Beta3 [125I]Iodopindolol ICI118551 Beta Binding (0.1 nM) Buffer

M1 [3H]QNB (0.5 nM) Atropine Muscarinic Binding Buffer

M2 [3H]QNB (0.5 nM) Atropine Muscarinic Binding Buffer

M3 [3H]QNB (0.5 nM) Atropine Muscarinic Binding Buffer

M4 [3H]QNB (0.5 nM) Atropine Muscarinic Binding Buffer

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M5 [3H]QNB (0.5 nM) Atropine Muscarinic Binding Buffer

CB1 [3H]CP55940 CP55940 Cannabinoid Binding Buffer

CB2 [3H]CP55940 CP55940 Cannabinoid Binding Buffer

Sigma1 [3H]Pentazocine Haloperidol Sigma Binding (3 nM) Buffer

Sigma2 [3H]DTG (3 nM) Haloperidol Sigma Binding Buffer

Raw dpm data from the Microbeta counter are analyzed on the PDSP DB. Total bound radioactivity is estimated from quadruplicate wells containing no test or reference compound and adjusted to 100%; non-specifically bound radioactivity is assessed from quadruplicate wells containing 10 μM of a suitable reference compound (see Table 1) and adjusted to 0%. The average bound radioactivity in the presence of the test compound (10 μM final assay concentration, quadruplicate determinations) is then expressed on the percent scale. The percent inhibition of radioligand binding is calculated as follows:

% inhibition = 100% - % radioactivity bound

The PDSP on-line data entry and analysis system calculates the variance of the quadruplicate determinations (for the total, non-specific, and test compound binding values) and variances greater than 20% are flagged for further inspection and assays are repeated if necessary. Additionally, % inhibition values that are greater than the total binding (i.e., 100%) by at least 20% are also flagged for inspection; such results could indicate allosteric modulation of radioligand binding.

Membrane Fraction Preparation

For binding assays using stably transfected cell lines, cells seeded (using 5% dialyzed fetal bovine serum-supplemented medium) in 10-cm dishes (90% confluent) are incubated overnight in either serum-free medium or medium containing 1% dialyzed fetal bovine serum. The next day, the cells are scraped into the medium and pelleted by centrifugation (1000 x g, 10 min). The cell pellet is resuspended in chilled 50 mM Tris, pH 7.4 (4 degrees) and triturated gently with a P1000 pipette tip to effect hypotonic lysis. The suspension is then centrifuged at 21,000 x g for 20 min to yield a crude membrane fraction pellet. The 50 mM Tris is removed and cells are either frozen on dry ice and maintained at -80 degrees centigrade until assay or resuspended immediately in a desired volume of appropriate assay buffer.

For binding assays using transiently transfected cell lines, cells are transfected as described in the Statement of Work. Twenty-four hours after transfection, the cells are split at an appropriate ratio (typically 1:2 or 1:3) into 10-

-298- cm dishes using 5% dialyzed fetal bovine serum-supplemented medium. Then, after overnight growth (generally 90% confluence is reached), the cells are incubated overnight in medium supplemented with no or 1% dialyzed fetal bovine serum. The next day (72 hours after transfection), the cells are scraped and processed as described for stable transfectants.

For binding assays using tissue, crude membrane fractions are prepared from rodent (typically rat) brain or kidney (purchased from PelFreeze Biologicals). Frozen tissue (maintained at -80 degrees centigrade) is thawed on ice, and then homogenized on ice in 10 vol of 50 mM Tris, pH 7.4 containing protease inhibitor cocktail (Roche) using a Polytron homogenizer (3 pulses, each of 10 sec). The homogenate is centrifuged at 40,000 x g for 20 min, and then the resulting supernatant is decanted and replaced with the same buffer. Two to three additional rounds of homogenization-centrifugation are performed to ensure thorough homogenization and also to wash away endogenous ligands (particularly important for GABA assays). The final, washed pellet is resuspended in the same buffer, homogenized one last time, and aliquoted into eppendorf tubes such that one tube contains sufficient material for 24 wells of a binding assay plate (generally 2.5-12.5 mg membrane protein/eppendorf tube, or 100 to 500 μg of membrane protein per well of the binding assay plate). The crude membrane fractions are stored at -80 degrees centigrade until assay, which in most instances is within 3 days of the crude membrane fraction preparation.

Experimental Procedure and Data Analysis for Secondary Screening of the Dopamine Transporter and the Beta3 Receptor

Dopamine Transporter Assay: Protocol adapted from Jensen et al. Neuropsychopharmacology 5 Dec (2007).

Assay Buffer: 1X Krebs-Ringer HEPES glucose buffer, 0.1 mM ascorbate, 0.1 mM pargyline, 0.1 mM tropolone (DAT and NET assays only), pH 7.4. Dopamine Transporter-expressing cell lines are seeded in 96-well, poly-L-lysine-coated plates 48 hours prior to the assay (40,000 cells per well) in DMEM containing 5% dialyzed serum. Twenty hours prior to the assay, the medium is changed to serum-free DMEM. On the day of the assay, the DMEM is washed and replaced with 30 μl of assay buffer containing vehicle or dilutions of test or reference compound (final concentrations ranging from 0.1 nM to 10 μM, each concentration assayed in triplicate). A 10-min pre-incubation is performed in a 37-degree centigrade, humidified incubator. Then, monoamine transport is initiated by addition of 30 μl of 2X [3H]monoamine, the specific activity of which is diluted 10- to 50-fold with non- labeled monoamine (final assay concentration of [3H]monoamine is 1 μM). Uptake of [3H]monoamine is allowed to proceed for 4 min, after which the buffer is aspirated and the cells are washed three times with ice-cold buffer containing 10 μM paroxetine (for SERT), GBR12909 (for DAT), or nisoxetine (for NET). Then, polystyrene-compatible scintillation cocktail (Microscint PS, PerkinElmer; 50 μl/well) is added and the plates are sealed and agitated on an orbital shaker at a high setting for 5 min. The plates are then counted in a Wallac MicroBeta TriLux scintillation counter.

Raw data (dpm) representing total [3H]monoamine uptake (i.e., specific + non- specific uptake) are plotted as a function of the logarithm of the molar concentration of the competitor (i.e., test or reference compound). Non-linear regression of the normalized (i.e., percent of [3H]monoamine uptake relative to that observed in the

-299- absence of test or reference compound) raw data is performed in Prism 4.0 using the built-in three parameter logistic model describing competitive inhibition (one-site):

y = bottom + [(top-bottom)/(1 + 10x-logIC50)] where bottom equals the best-fit non-specific [3H]monoamine uptake (i.e., non- specific uptake) and top equals the best-fit total [3H]monoamine uptake (i.e., uptake absent any competitor). The log IC50 (i.e., the log of the test or reference compound concentration that reduces [3H]monoamine uptake binding by 50%) is thus estimated from the data and used to obtain the Ki by applying the Cheng-Prusoff approximation: PI: Bryan L. Roth MD, PhD U n i v e r s i t y o f N o r t h C a r o l i n a a t C h a p e l H i l l S1.113 | National Institute of Mental Health Psychoactive Drug Screening Program

Ki = IC50/(1 + [monoamine]/KM) where [monoamine] equals the assay [3H]monoamine concentration and KM equals the affinity constant of the monoamine for the transporter.

Beta3 Receptor Assay: Protocol adapted from Jensen et al. Neuropsychopharmacology 5 Dec (2007).

Assay Buffer: 1X Krebs-Ringer bicarbonate glucose buffer, 0.75 mM IBMX, pH 7.4. Receptor-expressing cell lines are seeded in 96-well, poly-L-lysine-coated plates 48 hours prior to the assay (40,000 cells per well) in DMEM containing 5% dialyzed serum. Twenty hours prior to the assay, the medium is changed to serum-free DMEM. On the day of the assay, the DMEM is washed and replaced with 30 μl of assay buffer. A 10-min pre-incubation is performed in a 37-degree centigrade, humidified incubator. Then, the cells are stimulated by addition of 30 μl of 2X dilutions of test or reference compound (final concentrations ranging from 0.1 nM to 10 μM, each concentration assayed in triplicate). A positive control (100 μM forskolin) is also included. Accumulation of cAMP is allowed to continue for 15 min, after which the buffer is removed and the cells are lysed with Cell Lysis Buffer (CatchPoint cAMP Assay Kit, Molecular Devices). Next, the lysates are transferred to 96-well, glass-bottom plates coated with goat anti-rabbit IgG and adsorbed with rabbit anti-cAMP (Molecular Devices). Following a 5-min incubation, horseradish peroxidase-cAMP conjugate is added (Molecular Devices) and a 2-hour incubation is performed at room temperature. Then, after three washes with Wash Buffer (Molecular Devices), Stoplight Red substrate (Molecular Devices), reconstituted in Substrate Buffer (Molecular Devices) containing freshly-added 1 mM H2O2, is added and, after a 15-min incubation at room temperature, fluorescence is measured (excitation 510-545 nm, emission 565-625 nm). For each assay, a cAMP calibration curve is generated and controls without lysate and without antibody are included.

For agonist tests, raw data (maximum fluorescence, fluorescence units) for each concentration of test compound or reference agonist are normalized to the basal (vehicle-stimulated) fluorescence (reported as fold increase over basal) and plotted as a function of the logarithm of the molar concentration of the drug (i.e., test or reference compound). Non-linear regression of the normalized data is performed in Prism 4.0 (GraphPad Software) using the built-in three parameter logistic model

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(i.e., sigmoidal concentration-response) describing agonist-stimulated activation of one receptor population:

y = bottom + [(top-bottom)/(1 + 10x-logEC50)] where bottom equals the best-fit basal fluorescence and top equals the best-fit maximal fluorescence stimulated by the test compound or reference agonist. The log EC50 (i.e., the log of the drug concentration that increases fluorescence by 50% of the maximum fluorescence observed for the test compound or reference agonist) is thus estimated from the data, and the EC50 (agonist potency) is obtained. To obtain an estimate of the relative efficacy of the test compound (Rel. Emax), its best-fit top is compared to and expressed as a ratio of that for the reference agonist (Rel. Emax of the reference agonist is 1.00).

To ascertain whether test compounds are antagonists, a double-addition paradigm similar to that used in calcium mobilization assays is employed. First, 30 μl of test compound (20 μM) is added (10 μM final concentration) and a 15-min incubation is performed. Then, 30 μl of reference agonist (3X; EC90) is added (final concentration of agonist is EC30) and cAMP accumulation is allowed to proceed for 15 min. The samples are then processed for cAMP measurements as detailed above. Measurements of reference agonist-induced cAMP accumulation are compared to the signals elicited by the reference agonist following addition of vehicle instead of test compound and expressed as a ratio. ‘Hits’ (compounds that antagonize reference agonist-stimulated increases in baseline-normalized fluorescence by at least 50%) are then characterized by a modified Schild analysis.

For modified Schild analysis, a family of reference agonist concentration-response isotherms is generated in the absence and presence of graded concentrations of test compound (added 15 min prior to reference agonist). Theoretically, compounds that are competitive antagonists cause a dextral shift of agonist concentration-response isotherms without reducing the maximum response to agonist (i.e., surmountable antagonism). However, on occasion, factors such as non-competitive antagonism, hemiequilibria, and/or receptor reserve cause apparent insurmountable antagonism. To account for such deviations, we apply the modified Lew-Angus method to ascertain antagonist potency (Christopoulos et al., 1999). Briefly, equieffective concentrations of agonist (concentrations of agonist that elicit a response equal to the EC25% of the agonist control curve) are plotted as a function of the test compound concentration present in the wells in which they were measured. Non- linear regression of the baseline-normalized data is performed in Prism 4.0 using the following equation: pEC25% = -log ([B] + 10-pK) - log c where EC25% equals the concentration of agonist that elicits a response equal to 25% of the maximum agonist control curve response and [B] equals the antagonist concentration; K, c, and s are fit parameters. The parameter s is equal to the Schild slope factor. If s is not significantly different from unity, pK equals pKB; otherwise, pA2 is calculated (pA2 = pK/s). The parameter c equals the ratio EC25%/[B].

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Experimental Procedure for In Vivo Mouse Peritonitis Testing

Administration of vehicle in 4 mice, 1µg PGE2 in 3 mice, or 1µg AM7645 in 4 mice was given by intravenous injection. After 5 minutes, peritonitis was initiated in all mice by an intraperitoneal injection of 1mg zymosan A in 1mL of sterile saline. At 4 hours, mice were killed, and peritoneal exudates were harvested (5ml DPBS -/- without calcium and magnesium). Resident monocytes/microphages (MΦs) were identified and enumerated by flourescence-activated cell sorting (FACS) and light microscopy.

Procedure for In Vitro ECIS Assays of Recombinant EP Receptors

Ligand-receptor dependent changes in impedance were monitored using an Electrical Cell-Substrate Impedance Sensing System (ECIS; Applied Biophysics, Troy, NY). CHO cells were transfected with EP1, EP3, EP4 receptors or mock plasmid. Forty-eight hours later, cells were plated unto an 8-well ECIS chamber (8W10E+) at 0.2x106 cells/well for overnight. The next day, culture medium was replaced with serum-free medium and compounds were added to the chambers (100 nM each). ECIS acquisition of impedance was monitored for 20-30 min at 37 ºC.

Mock transfected CHO cells displayed Gq-like response when treated with PGE2.

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Experimental Procedures for PFC Immunological Studies

Spleen cells and sheep red blood cells (SRBCs) were placed in tissue culture wells for 5 days. Lymphocytes that respond to the SRBCs differentiate and secrete antibody to the red blood cells. The number of antibody secreting lymphocytes was quantitated by harvesting the cells and adding to them an excess of SRBCs plus guinea pig complement. The lymphocytes and red blood cells were placed in chambers formed by two glass slides held together by double-sided sticky tape. The lymphocytes and red blood cells form a single layered lawn, where the red bloods vastly outnumber the lymphocytes. If a lymphocyte is secreting antibody to the red blood cells, in the presence of complement, the red blood cells surrounding the lymphocyte lyse, forming a plaque that can be seen easily in a dissecting microscope. The number of plaques was counted to determine the number of plaque- forming lymphocytes (PFCs). Control cultures received no additive or received the vehicle in which the prostamides are dissolved. Experimental wells received prostamides at various doses. Results are expressed as a Suppression Index which is:

#PFCs in experimental wells/#PFCs in no treatment control wells

Viability of cultures was assessed in the flow cytometer using the LIVE/DEAD stain kit (Molecular Probes, Inc).

Experimental Procedures for MLR Immunological Studies

Spleen cells from two different mouse strains that are histoincompatable were mixed together in wells in culture. Cells of one strain had been pretreated with mitomycin C to prevent cell division. When the cells of the responder strain recognize the cells of the stimulator strain as foreign, they divide. Cell division is measured by uptake of 3[H]-thymidine. After incubation for 48 hours and then addition of the thymidine for another 18 hours, the cells were harvested and the amount of radioactivity was determined as counts per minute (cpm). Compounds which are immunosuppressive inhibit the responder cells from dividing and thus decrease the amount of radioactivity that is incorporated. Viability of cultures was assessed in the flow cytometer using the LIVE/DEAD stain kit (Molecular Probes, Inc).

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Appendix 3: Supplemental Synthetic Methods and Characterization for Compounds Presented in Chapter 5

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General Information:

All reactions were carried out in flame-dried glassware under argon atmosphere. Corey’s lactone benzoate aldehyde starting material was purchased from Cayman Chemical and used as supplied. All other reagents were purchased from Sigma-Aldrich and used as supplied. Tetrahydrofuran, dichloromethane, diethyl ether, and acetonitrile were obtained from a dry solvent system (alumina) and used without further drying. Unless otherwise noted, reactions were magnetically stirred and monitored by thin layer chromatography with Merck 250 µm silica gel 60 Å plates. Flash chromatography was performed using Biotage Isolera Flash Purification with Luknova 40-60 µm 60 Å silica gel columns. Yields refer to chromatographically and spectroscopically pure compounds, unless otherwise noted. 1H spectra were taken in CDCl3, at 400 or 500 MHz as indicated. 13C NMR spectra were also taken in CDCl3 at 100 MHz. Chemical shifts are reported in parts per million relative to TMS 1 13 ( H, δ 0.00) or CDCl3 ( C, δ 77.0). Data are reported as follows: chemical shift, multiplicity (s = singlet, d = doublet, t = triplet, q = quartet, quin = quintet, m = multiplet, br = broad), coupling constant, integration. Diastereomeric ratios were determined by 1H NMR (500 MHz) analysis of crude mixtures. High resolution mass- spectra were obtained on a Waters Q-TOF Ultima spectrometer at University of Illinois, SCS, Mass Spectrometry Laboratory.

Experimental Methods

General Procedure for Preparation of the Phosphonates

A solution of dimethyl methylphosphonate (3.0 mL, 28.1 mmol) in anhydrous THF (90 mL) was cooled to -78 ºC under argon. n-BuLi (2.5 M in hexane, 12.0 mL, 30.0 mmol) was added dropwise and stirring was continued at -78 ºC for 90 min. ε- caprolactone (1.5 mL, 13.5 mmol) (or γ-butyrolactone or methyldecanoate) was added and after 1 hr, oxalic acid (2.60g, 28.8mmol) was added and stirring was continued for 30 min warming to rt. The reaction mixture was filtered over Celite, washed with THF, and concentrated. Purification by flash chromatography on silica gel (MeOH/DCM, 5:95) afforded 3.16g (98%) of the desired product as a colorless oil.

Dimethyl 7-hydroxy-2-oxoheptylphosphonate

O MeO P OMe

OH O 1 H NMR (400MHz, CDCl3) δ [ppm] = 3.79 (d, JHP = 11.54 Hz, 6H), 3.62 (t, J = 6.53 Hz, 2H), 3.10 (d, JHP = 23.09 Hz, 2H), 2.64 (t, J = 7.28 Hz, 2H), 2.47 (br. s., 1H) + 1.66-1.53 (m, 4H), 1.41 – 1.35 (m, 2H). HRMS calcd for C9H20O5P, [M+H ] 239.1048, found 239.1043.

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Dimethyl 5-hydroxy-2-oxopentylphosphonate O MeO P OMe

OH O

1 H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.82 (dd, JHP = 20.02 Hz, 11.23 Hz, 6H), 3.68-3.60 (m, 2H) 3.13 (d, JHP = 22.46, 2H), 2.77 (t, J = 6.59Hz, 2H),1.87 (quin, J = 6.35 Hz, 2H)

Dimethyl 2-oxo-undecylphosphonate O MeO P OMe

O 1 H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.79 (d, JHP = 11.23 Hz, 6H), 3.08 (d, JHP = 22.46 Hz, 2H), 2.61 (t, J = 7.57 Hz, 2H), 1.58 (t, J = 6.84 Hz, 2H), 1.33-1.21 (m, 12H), 0.88 (t, J=6.84 Hz, 3 H)

General Procedure for Preparation of the Protected Phosphonates

To a solution of dimethyl 7-hydroxy-2-oxoheptylphosphonate (2.31 g, 9.68 mmol) in anhydrous DCM (60 mL) at 0ºC was added imidazole (1.36 g, 20.35 mmol) and tert-butylchlorodimethylsilane (3.03 g, 20.10 mmol). The reaction was allowed to stir overnight and slowly warm to rt. The mixture was quenched with saturated NH4Cl solution. The organic phase was washed with water and brine, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (Acetone/Hexanes, 80:20) afforded 3.34g (98%) of the desired product as a colorless oil.

Dimethyl 7-(tert-butyldimethylsilyloxy)-2-oxoheptylphosphonate

O MeO P OMe

OTBS O

1 H NMR (400MHz, CDCl3) δ [ppm] = 3.79 (d, JHP = 11.54 Hz, 6H), 3.60 (t, J = 6.53 Hz, 2H), 3.09 (d, JHP = 23.09 Hz, 2H), 2.62 (t, J = 7.28 Hz, 2H), 1.60 (quin, J = 7.53 Hz, 2H), 1.52 (quin, J = 7.03 Hz, 2H), 1.37 – 1.29 (m, 2H), 0.89 (s, 9H), 0.04 (s, + 6H). HRMS calcd for C15H34O5SiP, [M+H ] 353.1913, found 353.1905.

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Dimethyl 5-(tert-butyldimethylsilyloxy)-2-oxopentylphosphonate O MeO P OMe

OTBS O

1 H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.79 (d, JHP = 11.23 Hz, 6H), 3.62 (t, J = 5.86 Hz, 2H), 3.11 (d, JHP = 22.46 Hz, 2H), 2.69 (t, J = 7.32 Hz, 2H), 1.80 (quin, J = 6.59 Hz, 2H), 0.88 (s, 9H), 0.04 (s, 6H)

General Procedure for Preparation of the Enones

To a solution of NaH (60% dispersion in mineral oil, 112.5 mg, 2.82 mmol) in anhydrous THF (120 mL) at 0 ºC was added the phosphonate (892.1 mg, 2.53 mmol). The reaction was stirred at rt for 1hr after which it was cooled to 0 ºC and corey’s lactone benzoate aldehyde (696.3 mg, 2.539 mmol) was added. After 2.5 hrs at rt the reaction was quenched with glacial acetic acid (0.5 mL). The mixture was concentrated, diluted with EtOAc, and washed with water and brine. The organic layers were combined, dried over MgSO4, and concentrated. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 90:10) afforded 1.27g (100%) of the desired product as a colorless oil.

(3aR,4R,5R,6aS)-4-((E)-8-(tert-butyldimethylsilyloxy)-3-oxooct-1-enyl-2- oxohexahydro-2H-cyclopenta[b]furan-5-yl benzoate

O O

O O OTBS O

1 H NMR (400MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.20 Hz, 2H), 7.58 (t, J = 7.54 Hz, 1H), 7.45 (t, J = 7.54 Hz, 2H), 6.70 (dd, J = 15.77 Hz, 7.54 Hz, 1H), 6.24 (d, J = 15.77 Hz, 1H), 5.33 (q, J = 5.49 Hz, 1H), 5.11 (t, J = 4.80 Hz, 1H), 3.59 (t, J = 6.34 Hz, 2H), 2.97 – 2.86 (m, 3H), 2.63 (dt, J = 15.77 Hz, 6.51 Hz, 1H), 2.58 – 2.47 (m, 3H), 2.30 (ddd, J = 15.63, 3.91, 2.00 Hz, 1H), 1.61 (quin, J = 7.54 Hz, 2H), 1.52 (quin, J = 6.86 Hz, 2H), 1.39 – 1.30 (m, 2H), 0.89 (s, 9H), 0.04, (s, 6H). HRMS + calcd for C28H41O6Si, [M+H ] 501.2672, found 501.2677.

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(3aR,4R,5R,6aS)-4-((E)-6-(tert-butyldimethylsilyloxy)-3-oxohex-1-enyl-2- oxohexahydro-2H-cyclopenta[b]furan-5-yl benzoate

O O

O O OTBS O

1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 7.32 Hz, 2H), 7.58 (t, J = 7.32 Hz, 1H), 7.45 (t, J = 7.32 Hz, 2H), 6.69 (dd, J = 15.87, 7.57 Hz, 1H), 6.25 (d, J = 15.63 Hz, 1H), 5.32 (q, J = 5.37 Hz, 1H), 5.11 (t, J = 5.37 Hz, 1H), 3.62 (t, J = 6.10 Hz, 2H), 2.85 - 2.97 (m, 3H), 2.59 - 2.70 (m, 3H), 2.52 (dt, J = 16.6, 5.86 Hz 1H), 2.31 (dd, J = 15.63, 3.42 Hz, 1H), 1.82 (quin, J = 6.71 Hz, 2H), 0.87 (s, 9H), 0.02 (s, 6H)

(3aR,4R,5R,6aS)-4-(E)-3-oxododec-1-enyl-2-oxohexahydro-2H- cyclopenta[b]furan-5-yl benzoate

O O

O O O

1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 7.81 Hz, 2H), 7.58 (t, J = 7.32 Hz, 1H), 7.46 (t, J = 7.81 Hz, 2H), 6.68 (dd, J = 15.63, 7.32 Hz, 1H), 6.24 (d, J = 16.11 Hz, 1H), 5.33 (q, J = 5.37 Hz, 1H), 5.11 (t, J = 5.13 Hz, 1H), 2.87 - 2.95 (m, 3H), 2.63 (dt, J = 15.63, 6.59 Hz, 1H), 2.49 - 2.56 (m, 3H), 2.32 (dd, J = 15.63, 3.42 Hz, 1H), 1.55 - 1.64 (m, 2H), 1.19 - 1.33 (m, 12H), 0.88 (t, J=6.84 Hz, 3 H)

Procedure for Preparation of (3aR,4R,5R,6aS)-4-((E)-8-oxy-3-oxooct-1- enyl-2-oxohexahydro-2H-cyclopenta[b]furan-5-yl benzoate

O O

O O OH O

TBS protected enone (1.523 g, 3.04 mmol) was taken in 5 mL THF and diluted with 15 mL H2O followed by 30 mL AcOH. The reaction was stirred at 40°C for 3.5 hours and then quenched with 50mL saturated sodium bicarbonate. The mixture

-308- was concentrated, washed with saturated sodium bicarbonate, water, and brine. The organic layer were combined, dried over magnesium sulfate, and concentrated. Purification by flash chromatography (Acetone/Hexane, 50:50) yielded 1.0475 g (89%) of the desired product as a colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 8.30 Hz, 2H), 7.58 (t, J = 6.84 Hz, 1H), 7.46 (t, J = 7.32 Hz, 2H), 6.70 (dd, J = 15.63, 6.84 Hz, 1H), 6.24 (d, J = 15.63 Hz, 1H), 5.34 (d, J = 4.88 Hz, 1H), 5.11 (br. s., 1H), 3.64 (t, J = 6.35 Hz, 2H), 2.84 - 2.98 (m, 3H), 2.64 (dt, J = 15.63, 7.21 Hz, 1H), 2.48 - 2.59 (m, 3H), 2.32 (dd, J = 15.63, 4.39 Hz, 1H), 1.60 - 1.68 (m, 2H), 1.53 - 1.60 (m, 2H), 1.38 (quin, J = 7.69 Hz, 2H)

General Procedure for Preparation of the Esters

To a solution of primary alcohol (46.9 mg, 0.122 mmol), DMAP (10 mg, 0.082 mmol) and triethylamine (90 µL, 0.646 mmol) in DCM (2mL) was added actyl chloride (20 µL, 0.281 mmol) (or trimethylacetylchloride or benzoylchloride, 1- adamantanecarbonylchloride). The reaction mixture was stirred at rt for 24 hours. The reaction was quenched with saturated ammonium chloride and then washed with water and brine. The organic layers were combined, dried over magnesium chloride, and concentrated. Purification by flash chromatography (Acetone/Hexane, 30:70) afforded 29.0 mg (87%) of the desired product as a colorless oil.

(3aR,4R,5R,6aS)-4-((E)-8-acetoxy-3-oxooct-1-enyl)-2-oxohexahydro-2H- cyclopenta[b]furan-5-yl benzoate

O O

O

O O O O

1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 7.81 Hz, 2H), 7.59 (t, J = 7.32 Hz, 1H), 7.46 (t, J = 7.81 Hz, 2H), 6.70 (dd, J = 15.87, 7.08 Hz, 1H), 6.24 (d, J = 15.63 Hz, 1H), 5.34 (d, J = 4.88 Hz, 1H), 5.11 (br. s., 1H), 4.05 (t, J = 6.59 Hz, 2H), 2.85 - 2.96 (m, 3H), 2.64 (dt, J = 15.63, 6.35 Hz, 1H), 2.49 - 2.58 (m, 3H) 2.32 (dd, J = 16.36, 5.62 Hz, 1H), 2.04 (s, 3H), 1.63 (quin, J = 7.32 Hz, 4H), 1.36 (quin, J = 7.69 Hz, 2H)

-309-

(3aR,4R,5R,6aS)-2-oxo-4-((E)-3-oxo-8-(pivaloyloxy)oct-1-enyl)hexahydro- 2H-cyclopenta[b]furan-5-yl benzoate

O O

O

O O O O

1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 7.81 Hz, 2H), 7.58 (t, J = 7.81 Hz, 1H). 7.46 (t, J = 7.32 Hz, 2H), 6.70 (dd, J = 15.63, 7.32 Hz, 1H), 6.24 (d, J = 15.63 Hz, 1H), 5.33 (d, J = 4.88 Hz, 1H), 5.11 (br. s., 1H), 4.04 (t, J = 6.59 Hz, 2H), 2.86 - 2.96 (m, 3H), 2.64 (dt, J = 15.63, 6.84 Hz, 1H), 2.49 - 2.58 (m, 3H), 2.32 (dd, J = 15.63, 4.39 Hz, 1H), 1.59 - 1.69 (m, 4H), 1.31 - 1.41 (m, 2H), 1.19 (s, 9 H)

(E)-8-((3aR,4R,5R,6aS)-5-(benzoyloxy)-2-oxohexahydro-2H- cyclopenta[b]furan-4-yl)-6-oxooct-7-enyl benzoate O O

O

O O O O

1H NMR (500 MHz, CHLOROFORM-d) δ ppm 8.03 (d, J = 8.30 Hz, 2H), 7.99 (d, J = 8.30 Hz, 2 H), 7.61 - 7.53 (m, 2H), 7.40 - 7.48 (m, 4 H), 6.70 (dd, J = 15.87, 7.08 Hz, 1H) 6.24 (d, J = 16.11 Hz, 1H), 5.29 - 5.36 (m, 1H), 5.10 (t, J = 4.88 Hz, 1H), 4.31 (t, J = 6.35 Hz, 2H), 2.85 - 2.96 (m, 3H), 2.55 - 2.67 (m, 3H), 2.45 - 2.55 (m, 1H), 2.31 (dd, J = 15.38, 4.64 Hz, 1H), 1.78 (quin, J = 7.08 Hz, 2H) 1.69 (quin, J = 7.45 Hz, 2H), 1.46 (quin, J = 7.69 Hz, 2H)

(E)-8-((3aR,4R,5R,6aS)-5-(adamantoyloxy)-2-oxohexahydro-2H- cyclopenta[b]furan-4-yl)-6-oxooct-7-enyl benzoate

O O

O

O O O O

1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 7.32 Hz, 2H), 7.58 (t, J = 7.32 Hz, 1H), 7.46 (t, J = 7.81 Hz, 2H), 6.70 (dd, J = 16.11, 7.32 Hz, 1H), 6.24 (d, J = 15.63 Hz, 1H), 5.33 (q, J = 5.37 Hz, 1H), 5.11 (t, J = 4.88 Hz, 1H), 4.03 (t, J =

-310-

6.59 Hz, 2H), 2.86 - 2.99 (m, 3H), 2.64 (dt, J = 15.38, 6.47 Hz, 1H), 2.47 - 2.58 (m, 3H), 2.32 (dd, J = 15.63, 3.42 Hz, 1H), 2.01 (br. s., 3H), 1.88 (br. s., 6H), 1.67 - 1.77 (m, 6H), 1.58 - 1.67 (m, 4H), 1.36 (quin, J = 7.69 Hz, 2H)

General Procedure for Preparation of the Chiral 15-Hydroxyls Final Compounds

A solution of enone (1.27 g, 2.54 mmol) in MeOH (100mL) was cooled to 0 ºC. Cerium (III) chloride heptahydrate (1.418 g, 3.81mmol) and sodium borohydride (144 mg, 3.81mmol) were added sequentially and the mixture was stirred for 30 min. It was concentrated, diluted with EtOAc, washed with water and brine, dried over MgSO4, and concentrated a second time. Purification by flash chromatography on silica gel (EtOAc/Hexanes, 60:40) gave pure 15S and 15R isomers in 10% de for a total of 1.09 g (86%) [0.60 g (S-isomer) and 0.49 g (R-isomer)] as a colorless oil.

(3aR,4R,5R,6aS)-4-((R/S,E)-8-(tert-butyldimethylsilyloxy)-3-hydroxyoct-1- enyl)-2-oxohexhydro-2H-cyclopenta[b]furan-5-yl benzoate

O O O O

O O O OTBS O OTBS OH OH

1 Isomer S H NMR (400 MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.32 Hz, 2H), 7.57 (t, J = 7.32 Hz, 1H), 7.44 (t, J = 7.81 Hz, 2H), 5.65 (dd, J = 15.63 Hz, 6.10 Hz, 1H), 5.56 (dd, J = 15.38 Hz, 7.08 Hz, 1H), 5.23 (q, J = 5.86 Hz, 1H), 5.06 (t, J = 5.62 Hz, 1H), 4.08 (q, J = 6.10 Hz, 1H), 3.56 (t, J = 6.47 Hz, 2H), 2.91 – 2.77 (m, 2H), 2.74 (q, J = 6.35 Hz, 1H), 2.61 (dt, J = 15.38 Hz, 6.59 Hz, 1H), 2.52 (d, J = 16.6 Hz, 1H), 2.23 (dd, J = 15.38 Hz, 3.91 Hz, 1H), 1.70 (br. s., 1H) 1.55 – 1.40 (m, 4H), + 1.40 – 1.25 (m, 4H), 0.89 (s, 9H), 0.03 (s, 6H). HRMS calcd for C28H43O6Si, [M+H ] 1 503.2829, found 503.2816. Isomer R H NMR (400 MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.32 Hz, 2H), 7.56 (t, J = 7.32 Hz, 1H), 7.44 (t, J = 7.81 Hz, 2H), 5.66 (dd, J = 15.63 Hz, 5.62 Hz, 1H), 5.58 (dd, J = 15.38 Hz, 6.84 Hz, 1H), 5.24 (q, J = 5.62 Hz, 1H), 5.06 (t, J = 5.62 Hz, 1H), 4.09 (q, J = 5.86 Hz, 1H), 3.58 (t, J = 6.35 Hz, 2H), 2.91 – 2.78 (m, 2H), 2.74 (q, J = 6.35 Hz, 1H), 2.60 (dt, J = 15.38 Hz, 6.59 Hz, 1H), 2.51 (d, J = 16.6 Hz, 1H), 2.23 (dd, J = 15.38 Hz, 3.91 Hz, 1H), 1.77 (br. s., 1H) 1.55 – 1.45 (m, 4H), 1.40 – 1.25 (m, 4H), 0.89 (s, 9H), 0.04 (s, 6H). HRMS + calcd for C28H43O6Si, [M+H ] 503.2829, found 503.2826.

-311-

(3aR,4R,5R,6aS)-4-((R/S,E)-6-(tert-butyldimethylsilyloxy)-3-hydroxyhex- 1-enyl)-2-oxohexhydro-2H-cyclopenta[b]furan-5-yl benzoate

O O O O

O O O OTBS O OTBS OH OH

1 Isomer S H NMR (400 MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.32 Hz, 2H), 7.57 (t, J = 7.32 Hz, 1H), 7.44 (t, J = 7.81 Hz, 2H), 5.65 (dd, J = 15.63 Hz, 6.10 Hz, 1H), 5.56 (dd, J = 15.38 Hz, 7.08 Hz, 1H), 5.23 (q, J = 5.86 Hz, 1H), 5.06 (t, J = 5.62 Hz, 1H), 4.08 (q, J = 6.10 Hz, 1H), 3.56 (t, J = 6.47 Hz, 2H), 2.91 – 2.77 (m, 2H), 2.74 (q, J = 6.35 Hz, 1H), 2.61 (dt, J = 15.38 Hz, 6.59 Hz, 1H), 2.52 (d, J = 16.6 Hz, 1H), 2.23 (dd, J = 15.38 Hz, 3.91 Hz, 1H), 1.70 (br. s., 1H) 1.55 – 1.40 (m, 2H), + 1.40 – 1.25 (m, 2H), 0.89 (s, 9H), 0.03 (s, 6H). HRMS calcd for C28H43O6Si, [M+H ] 1 503.2829, found 503.2816. Isomer R H NMR (400 MHz, CDCl3) δ [ppm] = 7.99 (d, J = 7.32 Hz, 2H), 7.56 (t, J = 7.32 Hz, 1H), 7.44 (t, J = 7.81 Hz, 2H), 5.66 (dd, J = 15.63 Hz, 5.62 Hz, 1H), 5.58 (dd, J = 15.38 Hz, 6.84 Hz, 1H), 5.24 (q, J = 5.62 Hz, 1H), 5.06 (t, J = 5.62 Hz, 1H), 4.09 (q, J = 5.86 Hz, 1H), 3.58 (t, J = 6.35 Hz, 2H), 2.91 – 2.78 (m, 2H), 2.74 (q, J = 6.35 Hz, 1H), 2.60 (dt, J = 15.38 Hz, 6.59 Hz, 1H), 2.51 (d, J = 16.6 Hz, 1H), 2.23 (dd, J = 15.38 Hz, 3.91 Hz, 1H), 1.77 (br. s., 1H) 1.55 – 1.45 (m, 2H), 1.40 – 1.25 (m, 2H), 0.89 (s, 9H), 0.04 (s, 6H). HRMS + calcd for C28H43O6Si, [M+H ] 503.2829, found 503.2826.

(3aR,4R,5R,6aS)-4-((R/S,E)-3-hydroxydodec-1-enyl)-2-oxohexahydro-2H- cyclopenta[b]furan-5-yl benzoate

O O O O

O O O O OH OH

Isomer S 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 7.81 Hz, 2H), 7.58 (t, J = 7.32 Hz, 1H), 7.46 (t, J = 7.81 Hz, 2H), 6.68 (dd, J = 15.63, 7.32 Hz, 1H), 6.24 (d, J = 16.11 Hz, 1H), 5.33 (q, J = 5.37 Hz, 1H), 5.11 (t, J = 5.13 Hz, 1H), 2.87 - 2.95 (m, 3H), 2.63 (dt, J = 15.63, 6.59 Hz, 1H), 2.49 - 2.56 (m, 3H), 2.32 (dd, J = 15.63, 3.42 Hz, 1H), 1.55 - 1.64 (m, 2H), 1.19 - 1.33 (m, 12H), 0.88 (t, J=6.84 Hz, 3 H) Isomer R 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.99 (d, J = 7.81 Hz, 2H), 7.58 (t, J = 7.32 Hz, 1H), 7.46 (t, J = 7.81 Hz, 2H), 6.68 (dd, J = 15.63, 7.32 Hz, 1H), 6.24 (d, J = 16.11 Hz, 1H), 5.33 (q, J = 5.37 Hz, 1H), 5.11 (t, J = 5.13 Hz, 1H), 2.87 - 2.95 (m, 3H), 2.63 (dt, J = 15.63, 6.59 Hz, 1H), 2.49 - 2.56 (m, 3H), 2.32 (dd, J = 15.63, 3.42 Hz, 1H), 1.55 - 1.64 (m, 2H), 1.19 - 1.33 (m, 12H), 0.88 (t, J=6.84 Hz, 3 H)

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(3aR,4R,5R,6aS)-4-((R/S,E)-8-acetoxy-3-hydroxyoct-1-enyl)-2- oxohexahydro-2H-cyclopenta[b]furan-5-yl benzoate

O O O O O O O O O O O OH O OH

Isomer R1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.25 - 1.42 (m, 4 H) 1.44 - 1.54 (m, 2 H) 1.55 - 1.62 (m, 2 H) 2.04 (s, 3 H) 2.24 (dd, J=15.14, 4.39 Hz, 1 H) 2.51 (d, J=17.58 Hz, 1 H) 2.62 (dt, J=15.63, 6.84 Hz, 1 H) 2.74 (q, J=6.51 Hz, 1 H) 2.77 - 2.91 (m, 2 H) 4.03 (t, J=6.84 Hz, 2 H) 4.07 - 4.15 (m, 1 H) 5.07 (t, J=5.86 Hz, 1 H) 5.25 (q, J=5.86 Hz, 1 H) 5.59 (dd, J=15.63, 6.84 Hz, 1 H) 5.66 (dd, J=15.14, 5.37 Hz, 1 H) 7.45 (t, J=7.81 Hz, 2 H) 7.57 (t, J=7.81 Hz, 1 H) 8.00 (d, J=7.81 Hz, 2 H) Isomer S 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.22 - 1.37 (m, 4 H) 1.44 - 1.58 (m, 4 H) 2.04 (s, 3 H) 2.23 (dd, J=15.63, 4.88 Hz, 1 H) 2.52 (d, J=17.58 Hz, 1 H) 2.63 (dt, J=15.63, 6.35 Hz, 1 H) 2.74 (q, J=6.84 Hz, 1 H) 2.77 - 2.92 (m, 2 H) 4.00 (t, J=6.59 Hz, 2 H) 4.09 (br. s., 1 H) 5.07 (t, J=6.35 Hz, 1 H) 5.24 (q, J=5.86 Hz, 1 H) 5.57 (dd, J=15.14, 7.32 Hz, 1 H) 5.65 (dd, J=15.14, 5.86 Hz, 1 H) 7.45 (t, J=7.32 Hz, 2 H) 7.57 (t, J=7.32 Hz, 1 H) 8.00 (d, J=7.81 Hz, 2 H)

(3aR,4R,5R,6aS)-4-((R/S,E)-3-hydroxy-8-(pivaloyloxy)oct-1-enyl)-2- oxohexahydro-2H-cyclopenta[b]furan-5-yl benzoate

O O O O

O O

O O O O O O OH OH

Isomer R 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.19 (s, 9 H) 1.28 - 1.40 (m, 4 H) 1.46 - 1.54 (m, 2 H) 1.56 - 1.60 (m, 2 H) 2.24 (dd, J=15.38, 4.15 Hz, 1 H) 2.51 (d, J=17.58 Hz, 1 H) 2.62 (dt, J=15.50, 6.65 Hz, 1 H) 2.75 (q, J=6.35 Hz, 1 H) 2.78 - 2.91 (m, 2 H) 4.02 (t, J=6.59 Hz, 2 H) 4.10 (d, J=2.93 Hz, 1 H) 5.07 (t, J=5.86 Hz, 1 H) 5.25 (q, J=5.86 Hz, 1 H) 5.59 (dd, J=15.63, 7.32 Hz, 1 H) 5.66 (dd, J=15.14, 5.37 Hz, 1 H) 7.45 (t, J=7.81 Hz, 2 H) 7.57 (t, J=7.32 Hz, 1 H) 8.00 (d, J=7.32 Hz, 2 H) Isomer S 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.19 (s, 9 H) 1.23 - 1.37 (m, 4 H) 1.44 - 1.58 (m, 4 H) 2.23 (dd, J=15.14, 3.91 Hz, 1 H) 2.52 (d, J=17.58 Hz, 1 H) 2.62 (dt, J=15.14, 6.84 Hz, 1 H) 2.74 (q, J=6.84 Hz, 1 H) 2.77 - 2.92 (m, 2 H) 4.00 (t, J=6.59 Hz, 2 H) 4.08 (q, J=6.35 Hz, 1 H) 5.06 (t, J=6.35 Hz, 1 H) 5.24 (q, J=6.19 Hz, 1 H) 5.57 (dd, J=15.63, 7.81 Hz, 1 H) 5.65 (dd, J=15.63, 6.35 Hz, 1 H) 7.45 (t, J=7.81 Hz, 2 H) 7.57 (t, J=7.32 Hz, 1 H) 7.99 (d, J=7.32 Hz, 2 H)

-313-

(R/S,E)-8-((3aR,4R,5R,6aS)-5-(benzoyloxy)-2-oxohexahydro-2H- cyclopenta[b]furan-4-yl)-6-hydroxyoct-7-enyl benzoate

O O O O

O O

O O O O O O OH OH

Isomer R 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.29 - 1.54 (m, 6 H) 1.68 - 1.78 (m, 2 H) 2.23 (dd, J=15.63, 3.91 Hz, 1 H) 2.51 (d, J=17.09 Hz, 1 H) 2.61 (dt, J=15.14, 6.84 Hz, 1 H) 2.70 - 2.90 (m, 3 H) 4.29 (t, J=6.59 Hz, 2 H) 5.06 (t, J=7.81 Hz, 1 H) 5.25 (q, J=5.86 Hz, 1 H) 5.59 (dd, J=15.63, 7.32 Hz, 1 H) 5.66 (dd, J=15.14, 5.37 Hz, 1 H) 7.39 - 7.49 (m, 4 H) 7.53 - 7.61 (m, 2 H) 7.99 (d, J=7.32 Hz, 2 H) 8.04 (d, J=7.32 Hz, 2 H) Isomer S 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.34 - 1.43 (m, 4 H) 1.45 - 1.58 (m, 2 H) 1.63 - 1.74 (m, 2 H) 2.23 (dd, J=15.14, 4.39 Hz, 1 H) 2.52 (d, J=17.58 Hz, 1 H) 2.58 - 2.67 (m, 1 H) 2.69 - 2.92 (m, 3 H) 4.10 (d, J=5.86 Hz, 1 H) 4.27 (t, J=6.59 Hz, 2 H) 5.06 (t, J=5.86 Hz, 1 H) 5.24 (q, J=5.86 Hz, 1 H) 5.57 (dd, J=15.63, 7.81 Hz, 1 H) 5.65 (dd, J=15.63, 6.35 Hz, 1 H) 7.40 - 7.48 (m, 4 H) 7.52 - 7.59 (m, 2 H) 7.99 (d, J=7.81 Hz, 2 H) 8.03 (d, J=7.32 Hz, 2 H)

(R/S,E)-8-((3aR,4R,5R,6aS)-5-(adamantoyloxy)-2-oxohexahydro-2H- cyclopenta[b]furan-4-yl)-6-hydroxyoct-7-enyl benzoate

O O O O

O O

O O O O O O OH OH

Isomer R 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.25 - 1.41 (m, 4 H) 1.44 - 1.54 (m, 2 H) 1.53 - 1.60 (m, 2 H) 1.71 (td, J=12.21, 8.79 Hz, 6 H) 1.88 (br. s., 6 H) 2.01 (br. s., 3 H) 2.24 (m, J=16.60, 3.91 Hz, 1 H) 2.51 (d, J=17.58 Hz, 1 H) 2.62 (dt, J=15.63, 6.84 Hz, 1 H) 2.72 - 2.91 (m, 3 H) 4.01 (t, J=6.59 Hz, 2 H) 4.10 (d, J=2.93 Hz, 1 H) 5.07 (t, J=5.86 Hz, 1 H) 5.26 (q, J=5.86 Hz, 1 H) 5.59 (dd, J=15.63, 7.32 Hz, 1 H) 5.66 (dd, J=15.63, 5.86 Hz, 1 H) 7.45 (t, J=7.81 Hz, 2 H) 7.57 (t, J=7.32 Hz, 1 H) 8.00 (d, J=7.32 Hz, 2 H) Isomer S 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 1.26 - 1.36 (m, 4 H) 1.44 - 1.55 (m, 4 H) 1.71 (m, J=12.21, 12.21, 8.30 Hz, 6 H) 1.88 (br. s., 6 H) 2.01 (br. s., 3 H) 2.23 (dd, J=15.63, 3.91 Hz, 1 H) 2.52 (d, J=17.58 Hz, 1 H) 2.62 (m, J=15.63, 6.84, 6.84 Hz, 1 H) 2.72 - 2.91 (m, 3 H) 3.99 (t, J=6.35 Hz, 2 H) 4.05 - 4.11 (m, 1 H) 5.07 (t, J=6.35 Hz, 1 H) 5.24 (q, J=6.02 Hz, 1 H) 5.57 (dd, J=15.63, 7.32 Hz, 1 H) 5.65 (dd, J=15.63, 5.86 Hz, 1 H) 7.45 (t, J=7.81 Hz, 2 H) 7.57 (t, J=7.32 Hz, 1 H) 8.00 (d, J=7.32 Hz, 2 H)

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Appendix 4: Biological Experimental Procedures for Assays Presented in Chapter 5

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General Procedure for the rat FAAH 3 point fluorescent assay screening

To determine if prostamide intermediate compounds inhibit the metabolism of N- arachidonoyl, 7 amino-4-methyl coumarin amide (AAMCA) by rFAAH. The fluorescent assay was completed as follows:

Assay Buffer: (use MilliQ water) 50 mM HEPES 1 mM EDTA 0.1% BSA pH 7.4

Dilution Buffer: 50% DMSO 50% Assay Buffer

Assay: Each well in a plate (all black, flat bottom, 96 well Costar #3915) should have the following except for blanks and control

20 µL each compound dilution (final conc. of 1 µM, 10 µM and 100 µM) or reference FAAH prep to equal 15 µg protein (if protein conc is 3 mg/mL) Assay Buffer to make a final volume of 200 µL *Pre-incubate for 15 minutes at room temperature, shaking on the titer plate shaker* 2 µL of 1 mM AAMCA in DMSO (final conc. = 10 µM; 1x Km)

• The plate should be set up as described below. • In the blank, use Lysis Buffer (instead of enzyme) and AAMCA for 100% inhibition. • In the control, use Dilution Buffer (instead of compound dilution), AAMCA, enzyme for 0% inhibition. • The reference compound is AM-5206 (10 µM in DMSO) o 1.5 µM = 15 µL 10 µM in DMSO with 85 µL dilution buffer . Final = 150 nM o 150 nM = 3 µL 10 µM in DMSO with 197 µL dilution buffer . Final = 15 nM • No AMC standard curve is needed, as this is merely estimation and can be done in RFU. • Column 12 should be used to test compound’s fluorescent properties. • The plate should be put in the plate reader using the Gen5 protocol entitled “FAAH Three point Screen”. o Temperature = 25°C o Time = 4 hours o Kinetic read every 20 minutes, medium shaking for 10 seconds just prior to each read o Excitation = 360/40 nm o Emission = 460/40 nm

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Serial Dilutions:

Reaction Start with 10 mM Dilution Buffer concentration Compound in (µL) (µM) DMSO (µL) 100 10 90 10 10 90 1 10 90

Plate set-up: Low to high concentration (from left to right), if you change the order, then you may have to modify the order in ‘power exported’ excel output

1 2 3 4 5 6 7 8 9 10 11 12 A Blank CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 Control (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) Blank CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 Control B (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) C Blank CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 Control (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 D (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) E CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) Ref CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 Ref F Low (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) high (10 nM) (30 nM) G Ref CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 Ref Low (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) high (10 nM) (30 nM) Ref CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 Ref H Low (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) high (10 nM) (30 nM)

Reference Blank CMPD Control

5 µL Enz 5 µL Lys 5 µL Enz 5 µL Enz 20 µL AM- buff 20 µL cmpd 20 µL diln 5206 dilution 20 µL diln dilution buff 2 µL subste buff 2 µL subste 2 µL subste 173 µL assay 2 µL subste 173 µL 173 µL buffer 173 µL assay assay buffer assay buffer

Calculations: 1. Experiment will be autosaved into ‘D/program files/biotek/gen5/experiments/sasha’ as ‘FAAH Three Point Assay_date_time’. 2. Export the data to Excel by clicking “power export” and save the raw data. 3. Divide (the RFU3hr of the control minus the RFU3hr of each sample concentration) by the RFU3hr of the control to get an estimated % inhibition. Calculate the standard deviation between the triplicates and the stdt error. Repeat this assay if the std error is >5%. 4. If you get nearly 100% inhibitions for all the three concentrations, take it for three points in ‘nM’ concentrations instead of ‘µM’ concentrations. 5. Based on these three concentrations of compound and the estimated % inhibition, predict what the IC50 for this compound will be. If it is less than 50 µM, follow this assay up with an 8 point inhibition assay. 6. Control: FAAH (15 µg total protein) should hydrolyze 10 µM AAMCA at a rate of 1.5% per hour. This corresponds to 1000RFU = 0.1 µM AMC

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product and you should see maximum fluorescence in control wells as >3000 RFU.

General Procedure for the human FAAH 3 point fluorescent assay screening

To determine if prostamide intermediate compounds inhibit the metabolism of N- arachidonoyl, 7 amino-4-methyl coumarin amide (AAMCA) by hFAAH. The fluorescent assay was completed as follows:

Assay Buffer: (use MilliQ water) 50 mM HEPES 1 mM EDTA 0.1% BSA pH 7.4

Dilution Buffer: 50% DMSO 50% Assay Buffer

Assay: Each well in a plate (all black, flat bottom, 96 well Costar #3915) should have the following except for blanks and control

20 µL each compound dilution (final conc. of 1 µM, 10 µM and 100 µM) or reference __ µL FAAH prep to equal 15 µg protein __ µL Assay Buffer to make a final volume of 200 µL *Pre-incubate for 15 minutes at room temperature, shaking on the titer plate shaker* 2 µL of 1 mM AAMCA in DMSO (final conc. = 10 µM)

• The plate should be set up as described below. • In the blank, use Lysis Buffer (instead of enzyme) and AAMCA for 100% inhibition. • In the control, use Dilution Buffer (instead of compound dilution), AAMCA, enzyme for 0% inhibition. • The reference compound is AM-3506 (1 µM in DMSO) o 120 nM = 120 µL of 1 µM in DMSO with 880 µL dilution buffer . Final = 12 nM o 50 nM = 50 µL of 1 µM in DMSO with 950 µL dilution buffer . Final = 5 nM • No AMC standard curve is needed, as this is merely estimation and can be done in RFU. • Column 12 should be used to test compound’s fluorescent properties. • The plate should be put in the plate reader using the Gen5 protocol entitled “FAAH Three point Screen”. o Temperature = 25°C o Time = 4 hours o Kinetic read every 20 minutes, medium shaking for 10 seconds just prior to each read o Excitation = 360/40 nm o Emission = 460/40 nm

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Serial Dilutions:

Reaction Start with 10 mM Dilution Buffer concentration Compound in (µL) (µM) DMSO (µL) 100 10 90 10 10 90 1 10 90

Plate set-up: Low to high concentration (from left to right), if you change the order, then you may have to modify the order in ‘power exported’ excel output

1 2 3 4 5 6 7 8 9 10 11 12 Blank CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 CMPD #1 Control A (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) B Blank CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 CMPD #2 Control (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) C Blank CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 CMPD #3 Control (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 CMPD #4 D (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) E CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 CMPD #5 (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) Ref CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 CMPD #6 Ref F Low (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) high (5 nM) (12 nM) G Ref CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 CMPD #7 Ref Low (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) high (5 nM) (12 nM) Ref CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 CMPD #8 Ref H Low (1 µM) (1 µM) (1 µM) (10 µM) (10 µM) (10 µM) (100 µM) (100 µM) (100 µM) high (5 nM) (12 nM)

Reference Blank CMPD Control

5 µL Enz 5 µL Lys 5 µL Enz 5 µL Enz 20 µL AM- buff 20 µL cmpd 20 µL diln 5206 dilution 20 µL diln dilution buff 2 µL subste buff 2 µL subste 2 µL subste 173 µL assay 2 µL subste 173 µL 173 µL buffer 173 µL assay assay buffer assay buffer

Calculations: 7. Experiment will be autosaved into ‘D/program files/biotek/gen5/experiments as ‘FAAH Three Point Assay_date_time’. 8. Export the data to Excel by clicking “power export” and save the raw data. 9. Divide (the RFU3hr of the control minus the RFU3hr of each sample concentration) by the RFU3hr of the control to get an estimated % inhibition. Calculate the standard deviation between the triplicates and the stdt error. Repeat this assay if the std error is >5%. 10. If you get nearly 100% inhibitions for all the three concentrations, take it for three points in ‘nM’ concentrations instead of ‘µM’ concentrations. 11. Based on these three concentrations of compound and the estimated % inhibition, predict what the IC50 for this compound will be. If it is less than 1 µM, follow this assay up with an 8 point inhibition assay. 12. Control: you should see maximum fluorescence in control wells as >3000 RFU.

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