synthesis and biological evaluation of novel endocannabinoid ......the cannabinoid receptors,...

SYNTHESIS AND BIOLOGICAL EVALUATION OF NOVEL ENDOCANNABINOID PROBES, METABOLICALLY STABLE ANALOGS, AND N-ACYLETHANOLAMINE-

HYDROLYZING ACID AMIDASE INHIBITORS

A dissertation presented

by

Kyle Mark Whitten

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

October, 2012

2

SYNTHESIS AND BIOLOGICAL EVALUATION OF ENDOCANNABINOID PROBES,

METABOLICALLY STABLE ANALOGS, AND N-ACYLETHANOLAMINE-HYDROLYZING ACID AMIDASE INHIBITORS

By

Kyle Mark Whitten

ABSTRACT OF DISSERTATION

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

October, 2012

3

The design of novel selective ligands is important for the structural characterization of G-

protein coupled receptors (GPCR). For this reason we have sought to study the interactions of

selective cannabinergic ligands with the CB1 and CB2 cannabinoid receptors to obtain

information on the pharmacophoric requirements for ligand-drug activities.

Our laboratory has developed ligand-assisted protein structure (LAPS) an approach

which involves the combined use of covalent ligands, CB1 and CB2 mutants, as well as LC/MS

based proteomic methods. We have designed a convergent synthesis for a series of potent and

metabolically stable anandamide based covalent ligands. These ligands contain either an azide or

isothiocyanate group in strategic sites for covalent binding to determine where this class of

ligands activates the GPCR.

2-Arachidonoylglycerol (2-AG) is an important endogenous signaling molecule towards

the cannabinoid receptors, however, the chemical and metabolic instability of 2-AG is a

hindrance to its study. Previous chemical syntheses used conditions that promote acyl migration

to the more stable 1(3)-AG. Thus, we have developed an efficient and condition neutral

chemoenzymatic synthesis where minimal, if any, acyl migration is observed. Concordantly, in

vivo study of 2-AG is difficult due to its short duration of action caused by hydrolysis from

monoacylglycerol lipase (MAGL). Synthesis of a metabolically stable 2-AG analog has been

developed to prevent MAGL metabolism.

Palmitoylethanolamine (PEA) is recognized as a naturally occurring anti-inflammatory

agent that has been shown to enhance the effect of anandamide. The metabolism of PEA is

controlled by N-acylethanolamine-hydrolyzing acid amidase (NAAA) and inhibition of this

enzyme will increase PEA levels. Thus, we have synthesized a series of selective and reversible

inhibitors of NAAA with IC50 values in the low nM range.

4

To Agnes,

for the sacrifices she made

5

ACKNOWLEDGMENTS

I would first like to give gratitude to my advisor Professor Alexandros Makriyannis for

giving me the opportunity to conduct research on significant and novel projects relating to the

endocannabinoid system. His guidance and scientific input helped the evolution of these projects

where we were able to obtain new understanding of the cannabinoid system. It has been a

pleasure to learn from him not only in organic and medicinal chemistry, but also understand the

science at the interface of chemistry and biology.

I am grateful for the time and advice of my thesis committee, Dr. Graham Jones, Dr.

Carol Paronis, and Dr. Robert Hanson. Their input and discussion was helpful in completing this

dissertation.

I would like to specially thank my lab mentor Dr. Kumar Vadivel for his guidance at the

bench. His organic chemistry knowledge has been most useful in overcoming the various

hurdles I have approached throughout my research. His insight and direction has been most

helpful in the specific chemistries herein. I would also like to thank senior chemists in our lab,

Dr. Kiran Vemuri, and Dr. Spyros Nikas for their helpful discussions throughout the years and

their invaluable knowledge in organic synthesis.

I would like to thank Dr. Jodi Wood for her work in the biochemistry lab, making sure I

obtained all relevant biochemical data for my compounds. Han Zhou, who was responsible for

completing all the covalent assays on the endocannabinoid probes I have synthesized. Dr. Jay

West, was my biochemistry counterpart on the N-acylethanolamine-hydrolyzing acid amidase

(NAAA) project. He has done a great job purifying human NAAA and screening all the

inhibitors I produced in a high throughput screen that he developed. Dr. Anna Bowman, who

modeled my endocannabinoid covalent probes interacting with human CB2 receptor. Dr. Toby

6

Järbe and Roger Gifford, who conducted pharmacological testing on mice with my 2-AG

analogs. And Dr. Roger Kautz, who has been a great help with the NMR. I would also like to

thank all the CDD members past and present who I’ve had the privilege of knowing throughout

these five years.

I want to thank Dr. Travis Dudding, who gave me my first opportunity to conduct organic

synthesis research in his lab at Brock University. This opportunity opened my eyes to research

which I have been doing ever since. Also, my undergraduate advisor Dr. Eddie Luzik, who

helped grow my skills as an organic chemist and helped me obtain funding for an organic

research project.

I want to thank my good friends Dr. Rose Gathungu, Dr. Chris Allen, and Lenny Dao; we

all came into the program together and are all leaving in the same year. I am grateful for your

friendship and enjoyed eating lunch together almost every day.

Last but not least, I would like to thank my family; my Mom and Dad who have been so

supportive throughout my life. They have encouraged me to pursue my interests and were

always there for me with whatever help I needed. As well as my little brothers, Eric and Mark,

who growing up with helped me become the person I am today. And most importantly, I want to

thank my wife Agnes; who decided to move with me to Boston after we graduated together from

University of New Haven. She has sacrificed a lot, and I am so grateful for her love and support.

7

TABLE OF CONTENTS

Abstract 2

Acknowledgements 5

Table of Contents 7

List of Figures 11

List of Tables 13

List of Schemes 14

Abbreviations 16

Chapter 1: The Endocannabinoid System

1.1 History of Cannabis Sativa 20

1.2 The Cannabinoid Receptors 20

1.2.1 Cannabinoid Receptor Distribution 21

1.2.2 Cannabinoid Receptor Modulation 22

1.2.3 Compounds Interacting with the Cannabinoid Receptors 22

1.3 The Endocannabinoids 26

1.3.1 Endocannabinoid Biosynthesis 27

1.4 Metabolizing Enzymes 30

1.4.1 Fatty Acid Amide Hydrolase -- FAAH 30

1.4.2 Monoacylglycerol Lipase -- MGL 32

1.4.3 N-Acylethanolamine-Hydrolyzing Acid Amidae -- NAAA 33

1.5 Endocannabinoid System Retrograde Signaling 34

1.6 Goal of Thesis Research 34

1.7 References 36

8

Chapter 2: Novel Endocannabinoid Probes

2.1 Introduction 44

2.2 Current Cannabinoid Probes 45

2.3 Design of Bifunctional Endocannabinoid Covalent Probes 49

2.4 Linear synthetic strategy 50

2.5 Development of a Convergent Synthesis for Covalent Probes 51

2.6 Binding Affinity and Covalent Binding Data of Covalent Probes 54

2.7 Head-group Optimization 56

2.8 Ligand-Assisted Protein Structure (LAPS) 60

2.8.1 LAPS Studies Utilizing AM9017 61

2.9 Significance of Transmembrane Helix 6 66

2.10 Docking of AM9017 into hCB2 Models 66

2.11 Conclusions 70

2.12 Experimental 72

2.13 References 90

Chapter 3: Chemoenzymatic Synthesis of Biologically Active Compounds

3.1 Introduction 94

3.1.1 Synthesis of 2-AG 95

3.1.2 Lipases in the Synthesis of Acylated Glycerols 97

3.2 Synthesis of 2-MAGs with Immobilized Candida antarctica and Rhizomucor

miehei

98

3.2.1 Synthesis of 2-MAGs with Immobilized Candida antarctica 98

9

3.2.2 Synthesis of 2-MAGs with Rhizomucor miehei 100

3.3 Enzymatic synthesis of N-Acylethanolamines 102

3.3.1 Candida antarctica for the direct aminolysis of esters 103

3.3.2 Reaction optimization and results 104

3.4 Conclusions 107


3.6 References 125

Chapter 4: Chemically and Metabolically Stable 2-Arachidonoylglycerol Analogs

4.1 Introduction 130

4.2 Synthesis of Biphenyl 2-AG Analogs 132

4.2.1 Biphenyl 2-AG Analog Cannabinoid Binding Data 137

4.3 Modification of the Glycerol Head Group 140

4.3.1 Synthesis of 2,4-dihydroxypentan-3-arachidonoates 140

4.3.2 Synthesis of 1,3-dihydroxybutan-2-arachidonoates 142

4.4 Purification and Isolation of 2,4-dihydroxypentan-3-arachidonoate Isomers 144

4.5 Identification of pent-2,4-ol-3-arachidonoate Stereochemistry 147

4.6 Chemically and Metabolically Stable 2-AG Analog Data 151

4.6.1 Chemical Stability of 2-AG Analogs 151

4.6.2 Metabolic Stability of 2-AG Analogs Compared to 2-AG 152

4.6.3 Cannabinoid In Vitro Binding Assay Data 154

4.6.4 Pharmacological Data of 2-AG, 1-AG, AA, and 2-AG Analogs In Vivo 155

4.7 Conclusions 159


10

4.9 References

180

Chapter 5: N-Acylethanolamine-Hydrolyzing Acid Amidase Inhibitors

5.1 Introduction and Background 183

5.1.1 Current NAAA inhibitors 184

5.2 Design, Synthesis, and Biological Evaluation of NAAA Inhibitors 186

5.2.1 Retroamides 186

5.2.2 Carbonates and Carbamates 188

5.2.3 Optimization of AM9058 192

5.2.4 Isothiocyanates 193

5.3 Evaluation of AM9053 Mode of Inhibition 198

5.4 Conclusions 202


5.6 References 227

Chapter 6: Future Directions

6.1 Novel Endocannabinoid Probes 231

6.2 Chemoenzymatic Methods 232

6.3 2-AG Analogs 232

6.4 NAAA Inhibitors 232

6.5 References 233

Appendix I : Publications 234

11

LIST OF FIGURES Figure 1.1 Δ9-Tetrahydrocannabinol 20

Figure 1.2 Class examples of compounds activating CB1 and CB2 receptors 23

Figure 1.3 Selective CB1 Agonists 24

Figure 1.4 CB1 receptor antagonists and inverse agonists 25

Figure 1.5 CB2 selective agonists 25

Figure 1.6 CB2 selective inverse agonists/antagonists 26

Figure 1.7 The primary endocannabinoids Anandamide and 2-Arachidonoyl glycerol 26

Figure 1.8 Other possible endocannabinoids 27

Figure 1.9 FAAH inhibitors 32

Figure 1.10 MGL inhibitor JZL184 33

Figure 2.1 THC based covalent probes 45

Figure 2.2 Other endocannabinoid probes 46

Figure 2.3 Tagged endocannabinoid probes 47

Figure 2.4 Arachidonoylcyclopropylamine covalent probes 48

Figure 2.5 CB1 receptor with transmembrane cysteines highlighted 60

Figure 2.6 Human CB2 receptor with transmembrane cysteines highlighted 61

Figure 2.7 Covalent assay of AM9017 with WT hCB2 and mCB2 63

Figure 2.8 Covalent assay of AM9017 with C6.47S hCB2 and mCB2 mutants 64

Figure 2.9 AM9017-hCB2 covalent complex equilibrated in a POPC membrane 66

Figure 2.10 Zoom-in of AM9017 covalent attachment to the hCB2 receptor 67

Figure 2.11 Comparison of inactive hCB2 (orange) with active hCB2 receptor

covalently bound to AM9017 (green) 68

12

Figure 3.1 N-Acylethanolamines 101

Figure 4.1 2-AG and published 2-AG analogs 130

Figure 4.2 1H NMR of 1,3-diol products after NaBH4 reduction of 1,3-diketone 144

Figure 4.3 Product separation from TLC analysis of tested solvent systems 146

Figure 4.4 Theoretical products from NaBH4 reduction of a 2-4 diketone 147

Figure 4.5 Zoom in on splitting patterns of C2, C3, and C4 protons 148

Figure 4.6 Newman projects of the two meso compounds: 49 & 50 150

Figure 4.7 Locomotor activity following administration of 2-AG analogs 155

Figure 4.8 Rearing activity following administration of 2-AG analogs 156

Figure 4.9 Locomotion and rearing of dosed mice with 2-AG, 1-AG, and arachidonic

acid 157

Figure 5.1 Best NAAA inhibitors derived from palmitic esters, 1 & 5; retroesters, 2;

retroamides, 3; along with palmitoylethanolamine (PEA) 4 184

Figure 5.2 Reported β-lactone NAAA inhibitors 185

Figure 5.3 Concentration dependent inhibition of human NAAA by AM9053 199

Figure 5.4 Tryptic digest of purified human NAAA obtained by MALDI-TOF MS for

protein neat (A) and AM9053 treated enzyme (B) 200

Figure 5.5 Lineweaver-Burk plot analysis of AM9053 inhibition of hNAAA 201

13

LIST OF TABLES Table 2.1 Binding affinity (Ki) for ligands 17-24 to the cannabinoid receptors 53

Table 2.2 Percentage of occupied receptors for 18-20 and 22-24 in covalent binding

assay 54

Table 2.3 Binding affinity (Ki) for ligands 62-65 to the cannabinoid receptors 58

Table 2.4 Percentage of occupied receptors by 62-65 in covalent binding assays 58

Table 3.1 Structure and Yields of Lipase catalyzed 2-MAGs 100

Table 3.2 Amidation of esters with immobilized Candida antarctica in 1:1 hexane-

diisopropylether 104

Table 4.1 2-point cannabinoid receptor binding assay data 137

Table 4.2 hMGL substrate assay results 152

Table 4.3 Cyclic AMP assay for 2-AG dimethyl analogs 154

Table 5.1 IC50 values for compounds 12-20 towards NAAA, FAAH, and MGL

enzymes 187

Table 5.2 Carbonates and Carbamates 190

Table 5.3 Carbamate inhibitors with alternate phenyl leaving groups 192

Table 5.4 Inhibition data of isothiocyanate inhibitors 196

14

LIST OF SCHEMES Scheme 1.1 N-acylethanolamine biosynthetic pathway 28

Scheme 1.2 Other possible biosynthetic routes of NAEs 29

Scheme 1.3 Biosynthetic pathways of 2-AG 30

Scheme 2.1 Possible mechanism of nitrene formation and nucleophilic attack of

isothiocyanate 44

Scheme 2.2 Design of bifunctional azide and isothiocyanate covalent probes 49

Scheme 2.3 Linear synthesis of 20-hydroxy methyl tetraynoate 49

Scheme 2.4 Convergent synthesis of tetraeneoates 51

Scheme 2.5 Deprotection of 10-(trimethylsilyl)deca-6,9-diyn-1-ol 51

Scheme 2.6 Functionalizing the arachidonoate tail 52

Scheme 2.7 Crude visualization of linear probes degree of freedom 55

Scheme 2.8 Anandamide and AM356 CB1 binding 56

Scheme 2.9 Synthesis of chiral head group 56

Scheme 2.10 Metabolically stable covalent probes 57

Scheme 3.1 Acyl migration from 2-AG to 1(3)-AG 93

Scheme 3.2 TIPS method for synthesizing 2-AG 94

Scheme 3.3 2-AG synthesis through benzylidene protected glycerol 95

Scheme 3.4 2-AG synthesis through glycidal ring openeing 95

Scheme 3.5: “AAA” and “ABA” triglycerides 96

Scheme 3.6 Chemoenzymatic Syntheses with immobilized Candida antarctica and

Rhizomucor miehei 98

Scheme 3.7 CAL catalyzed aminolysis of esters 104

15

Scheme 4.1 Synthesis of biphenyl 2-AG analogs 132

Scheme 4.2 Continued synthesis of biphenyl 2-AG analogs 134

Scheme 4.3 Synthesis of biphenyl ether 2-AG analogs 135

Scheme 4.4 Synthesis of to 2,4-dihydroxypentan-3-arachidonoates 140

Scheme 4.5 Attempted keto-oxirane ring opening 141

Scheme 4.6 Chemoenzymatic esterification of ±butane-1,2,3-triol 142

Scheme 4.7 Successful synthetic strategy towards a monomethyl 2-AG analog 143

Scheme 5.1 Synthesis of retroamide inhibitors 186

Scheme 5.2 Synthesis of Carbonates 25-27 188

Scheme 5.3 Synthesis of Carbamates 33-36, 38 and 40-41 189

Scheme 5.4 Parallel synthesis of carbamates 191

Scheme 5.5 Synthesis of Isothiocyanate NAAA inhibitors 194

Scheme 5.6 S-alkylation inhibition mechanism of β-lactones 198

Scheme 5.7 Irreversible inhibition by AM6701 198

16

ABBREVIATIONS

1,3-DBG 1,3-dibutoyl glycerol

2-AG 2-arachidonoyl glycerol

2-AGE 2-arachidonoyl glycerol ether

2-MAG 2-monoacylglycerol

AA arachidonic acid

Abh4 α/β-hydrolase-4

ACEA arachidonoyl 2'-chloroethylamine

ACPA arachidonoyl cyclopropylamine

AD Alzheimer's disease

AEA arachidonoyl ethanolamine

BOC tert-butoxycarbonyl

CAL Candida antarctica lipase

cAMP cyclic adenosine monophosphate

CB1 cannabinoid receptor 1

CB2 cannabinoid receptor 2

CNS central nervous system

DAG diacylglycerol

DBU 1,8-diazabicyclo[5.4.0]undec-7-ene

DGL diacylglycerol lipase

DMAP 4-dimethylamino pyridine

17

DPPA diphenylphosphoroyl azide

EC endocannabinoid

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

FAAH fatty acid amide hydrolase

GPCR G-protein couped receptor

GP-NAE glycerophospho-NAE

hCB human cannabinoid receptor

IC50 half maximal inhibitory concentration

Ki binding affinity

LAPS Ligand-assisted Protein Structure

LCMS liquid chromatography mass spectrometry

MALDI-TOF matrix assisted laser desorption/ionization - time of flight

MAP mitogen-activated protein

mCB mouse cannabinoid receptor

MGL monoacylglycerol lipase

NAAA N-acylethanolamine-hydrolyzing acid amidase

NAE N-acylethanolamine

NAE-P NAE-phosphate

NAPE N-acylphosphatidylethanolamine

NAPE-PLD NAPE-hydrolyzing phospholipase D

NAT N-acyl transferase

NMR nuclear magnetic resonance

OEA oleoylethanolamine

18

PA phosphatidic acid

PAMCA N-(4-methyl coumarin) palmitamide

PC phosphatidylcholine

PE phosphatidylethanolamine

PEA palmitoylethanolamine

PI phosphatidylinositol

PLA1 phospholipase A1

PMSF phenylmethylsulfonyl flouride

POPC palmitoyloleoylphosphatidylcholine

PPAR-α peroxisome proliferator-activated receptor-α

rCB rat cannabinoid receptor

SAR structure-activity relationship

TBAF tetrabutlyammonium fluoride

TFA trifluoroacetic acid

THC tetrahydrocannabinol

THF tetrahydrofuran

TIPS triisopropylsilyl

TLC thin layer chromatography

TMH transmembrane helix

TMS trimethylsilyl

UV ultraviolet

WT wild type

19

CHAPTER 1

THE ENDOCANNABINOID SYSTEM

20

1.1 History of Cannabis sativa

Cannabis sativa, commonly known as marijuana, has historically been used for its many

beneficial properties. Cannabis has been used since c. 2700 BC for its value in food, fiber, oil,

paper, and maybe most importantly—medicine. The Chinese were among the first to understand

the properties of marijuana consumption including their notes on psychoactive properties,

anesthesia, antiemetic, antibiotic, anthelmintic, as well as using it for over 100 additional

ailments.1 The primary psychoactive constituent responsible for many of the effects of cannabis

remained elusive until Δ9-tetrahydrocannabinol (THC, Figure 1.1) was eventually isolated and

characterized in 1964 by Mechoulam et al.2

O

OH

H

H

THC

Figure 1.1 Δ9-Tetrahydrocannabinol

1.2 The Cannabinoid Receptors

With the identification of THC, the next challenge was to determine how the body responds

when activated by cannabinoids and where this activation occurs in the body. Eventually, two

G-protein coupled receptors (GPCR) were identified, the cannabinoid 1 (CB1) receptor,

discovered in 19883 and cloned in 1990,4 and the cannabinoid 2 receptor (CB2), cloned in 1993.5

These receptors were first named as those of which respond to cannabinoids instead of naming

them after their endogenous ligands because there were no known endogenous agonists at the

time of their discovery.6 GPCRs are comprised of seven transmembrane helices connected by

seven intra and extracellular loops. The GPCRs can be divided into 5 main families: the

21

rhodopsin, adhesion, frizzled/taste, glutamate, and secretin families.7 GPCRs compromise the

largest family of pharmaceutical targets for modulation.

Between the CB1 and CB2 receptors is an overall 44% homology, with 68% amino acid

sequence homology between the transmembrane domains.5 Both receptors are coupled to Gi/o

(heterotrimeric) proteins, thus there activation adenylyl cyclase, and promote mitogen-activated

protein (MAP) kinase. This reduces the production of cyclic adenosine monophosphate

(cAMP).8

GPCRs are proposed to exist in an equilibrium between various activated states. There are

believed to be the active states (R’’, R*), inactive states (R’, R), and also the signaling state

(R*G). Various levels of basal activity can suggest which state the receptor models in a ligand-

free environment.9 Interactions with different types of ligands can shift the receptors equilibrium

to different states. Agonists are ligands that activate the receptor; they can be a full agonist in

where full efficacy is observed, or a partial agonist where efficacy is reduced compared to a full

agonist. Antagonists block the activation of the receptor by agonists and thus prevent or limit a

biological response from activation. Lastly, an inverse agonist can produce a negative cellular

response through lowering the activity below the basal level.10

1.2.1 Cannabinoid Receptor Distribution

The CB1 receptor primarily resides in the central nervous system (CNS).11 CB1 is one of

the most abundant GPCRs in the brain, and is predominately located on presynaptic terminals of

neurons,12-14 but they have also been discovered on postsynaptic neurons and on glial cells.15

Outside of the CNS, the CB1 receptor has been found in the gastrointestinal tract,16

cardiovascular system,17, 18 and reproductive system.19 The CB1 receptor is highly concentrated

in the brain, with dense accumulation in basal ganglia, cerebellum, and hippocampus which

22

affect motor control, cognitive function, and stress response. Much lower levels of CB1 are

observed in the lower brain stem.20

The CB2 receptor, however, is not commonly observed in the CNS,21 and is customarily

expressed in the immune system.22 The concentrations of the CB2 receptor were first measured in

the spleen, thymus, tonsils, and peripheral blood mononuclear cells.23 More recently, expression

of CB2 receptors in the spinal cord under neuropathic conditions has been discussed.24

1.2.2 Cannabinoid Receptor Modulation

CB receptor activation involves the inhibition of adenyl cyclase which leads to decreased

production of cAMP.25 Calcium influx and subsequent neurotransmitter release can also be

inhibited by CB receptor stimulation,26 while activation of ATP-sensitive potassium channels is

observed through the induction of antinociception,27 as well as MAP kinase activation.28, 29

Cannabinoid agonists have shown to activate subtypes of Gi/o α subunits by using a GTP

photoaffinity analog and measured through [35S]GTPγS binding.30 While mu and delta opioid

receptors can activate as many as 20 G-proteins, CB1 receptor binding has been shown to

activate only three G-proteins.31 [35S]GTPγS binding is utilized to measure agonist efficacy with

regards to G-protein activation.32

1.2.3 Compounds Interacting with the Cannabinoid Receptors

There are four main classes of compounds that are known to activate both the CB1 and

CB2 receptor. They include classical, nonclassical, aminoalkylindole, and eicosanoid (Figure

1.2). Classical cannabinoids are either plant derived or analogs of the tricyclic dibenzopyran

structure.33 Nonclassical compounds do resemble THC, however, they usually lack a pyran ring.

Pfizer developed CP55,940 which is one of the most widely used ligands for the study of the

cannabinoid system.34 WIN55,212-2 was one of the first compounds to produce pharmacological

23

effects similar to that of THC, in which the structure bare no resemblance to that of any classical

cannabinoid or nonclassical analog.35 Eicosanoids, represented by anandamide, are the class of

compounds which are biosynthesized on demand and referred to as endocannabinoids (EC),

which are discussed in Section 1.3.

Classical Nonclassical

O

OH

H

H

THC HO

OH

CP55,940

Aminoalkylindole Eicosanoid

O

NO

N

O

WIN55,212-2

O

NH

OH

Anandamide

Figure 1.2 Class examples of compounds activating CB1 and CB2 receptors

Selective CB1 agonists were developed to improve on the low selectivity of anandamide

through modification of the amide head group. Two of the more potent CB1 selective agonists are

arachidonoyl 2’-chloroethylamide (ACEA) and arachidonoyl cyclopropylamine (ACPA, Figure

1.3).36 ACEA has binding affinities of 1.4 nM and >2000 nM for CB1 and CB2 respectively

while ACPA has affinities of 1.4 nM and 715 nM respectively. However, these compounds lack

24

enzymatic stability towards fatty acid amide hydrolase (FAAH). This lead to the development of

(R)-methanandamide (see Chapter 2, Scheme 2.8) and O-1812, where the insertion of a 1’

methyl provided resistance to hydrolysis from FAAH.37

O

NH

Cl

ACEA

O

NH

ACPA

O

NH

OH

O-1812 CN

Figure 1.3 Selective CB1 Agonists

Examples of CB1 receptor antagonists and inverse agonists can be seen in Figure 1.4.

SR141716A, also known as Rimonabant, is a ligand developed by Sanofi-Aventis. First

classified as a CB1 selective antagonist with binding affinity of 1.98 nM for the CB1 receptor and

>1000 nM towards CB2,38 there is evidence it may also behave as an inverse agonist.39 More

recent reports indicate this inverse agonism may not be CB1 mediated.40 AM251 is a close analog

of Rimonabant with similar properties.41 LY320135 is chemically distinct from Rimonabant and

AM251, as it features a benzopyran as opposed to the diarylpyrazole of the other ligands, but it is

less potent towards the CB1 receptor with an affinity of 141 nM.42

25

NN

ONH

Cl

Cl

Cl

NN

ONH

Cl

Cl

I

SR141716A AM251

OO

O

O

CN

LY320135

Figure 1.4 CB1 receptor antagonists and inverse agonists

CB2 selective agonists can include a variety of structural motifs, including the classical

cannabinoid analog JWH133 (Figure 1.5) which exhibits CB1 and CB2 binding affinities of 677

nM and 3.4 nM respectively. AM1241 is an indole-based ligand with receptor affinities of 280

nM and 3.4 nM. Another selective agonist is the nonclassical-like HU308 with an affinity

towards CB2 of 22.7 nM. Potent CB2 selective inverse agonists/antagonists include SR144528

and AM630. The first is a diarylpyrazole and the later is an N-alkyl indole (Figure 1.6).

OH

H

JWH133

O

HU308

O

HO

N

O

N

NO2

IAM241

Figure 1.5 CB2 selective agonists

26

NN

ONH

Cl

SR144528

N

N

O

O

OI

AM630

Figure 1.6 CB2 selective inverse agonists/antagonists

1.3 The Endocannabinoids

An endocannabinoid, is an endogenous compound that interacts with the cannabinoid (CB)

receptors as its primary function.43 The two most studied endocannabinoids are anandamide

(AEA)44 and 2-arachiconoylglycerol (2-AG) (Figure 1.7).45 Additional endogenous CB agonists

include homo-γ-linolenoylethanolamide and docosatetraenoylethanolamide,46 noladin ether,47 N-

oleoyl dopamine48, N-arachidonoyl dopamine,49 oleamide and virodhamine (Figure 1.8).50

O

NH

OH

AEA

O

OOH

OH

2-AG

Figure 1.7 The primary endocannabinoids Anandamide and 2-Arachidonoyl glycerol

27

NH

OOH

docosatetraenoylethanolamide

NH

OOH

homo-y-linolenoylethanolamide

OOH

OH

noladin ether

NH2

O

oleamide

NH

OOH

OH

N-oleoyl dopamine

O

NH

N-arachidonoyl dopamine

OH

OH

O

ONH2

virodhamine

Figure 1.8 Other possible endocannabinoids

The first endocannabinoid was isolated from porcine brain in 1992. It was determined

that the structure was N-arachidonoylethanolamine (AEA), named anandamide from the Sanskrit

‘ananda’ meaning bliss and ‘amide’ for its chemical features .44 A second endocannabinoid, 2-

AG, was isolated from canine gut in 1995 and found to bind to the CB receptors.45

1.3.1 Endocannabinoid Biosynthesis

The hydrophobic properties of endocannabinoids do not allow them to be stored, rather,

they are biosynthesized based on necessity.51 Anadamide, belonging to the class of N-

acylethanolamines (NAEs), is formed in animal tissue through a common pathway. NAE

biosynthesis is commonly referred as the ‘transacylation-phosphodiesterase pathway’—where,

starting from glycerophospholipids, N-acyl transferase (NAT), and N-

acylphosphatidylethanolamine-hydrolyzing phospholipase D (NAPE-PLD) catalyze the two

steps which complete the process (Scheme 1.1).52,53,54

The transfer of a fatty acyl chain from the sn-1 position of glycerophospholipids (in

phosphatidylcholine [PC]) to the amino group of phosphatidylethanolamine (PE) is catalyzed by

the calcium dependent NAT to form N-acylphophatidylethanolamine (NAPE, Scheme 1.1). The

28

dependency of NAT on Ca2+ makes this first step the rate-limiting step.52, 53 In the second step,

NAPE-PLD hydrolyzes NAPE to NAE and phosphatidic acid (PA). 54

Scheme 1.1 N-acylethanolamine biosynthetic pathways

O

R1O

O R2

OO

POH

O

OH2N

PE

O

R3O

O R4

OO

POH

O

ON

PC

O

R1O

O R2

OO

POH

O

OHNR3

O NAPE

HO

O R4

OO

POH

O

ON

Lyso PC

NAT

OHHNR3

O

O

R1O

O R2

OO

POH

O

HOPANAE

NAPE-PLD

H2O

There are possible alternative pathways by which NAEs can be formed (Scheme 1.2).

Starting from NAPEs, hydrolysis of the O-acyl chains leads to either NAE-P (N-

acylethanolamine-phosphate) or lyso-NAPE. The phosphate group can be hydrolyzed from

NAE-P or the phosphodiester can be hydrolyzed from lyso-NAPE to form NAEs. Additionally,

α/β-hydrolase-4 (Abh4) can deacylate lyso-NAPE to glycerophospho-NAE (GP-NAE) which can

29

then be hydrolyzed to NAEs.54, 55 These alternative pathways may be just as important as NAPE-

PLD in the formation of NAEs. Cravatt et al. used NAPE-PLD(-/-) mice to measure NAPE-PLD

activity and endogenous levels of NAPEs and NAEs. As expected, NAPE-PLD activity was

decreased as well as saturated NAEs, however, the levels of polyunsaturated NAEs, including

anandamide, in the NAPE-PLD(-/-) mice were unchanged. This led to the conclusion that

NAPE-PLD is not the only contributor to the biosynthesis of NAEs.56

Scheme 1.2 Other possible biosynthetic routes of NAEs

O

R1O

O R2

OO

POH

O

OHNR3

O NAPE

OHP

OH

O

OHNR3

O

O

R1O

OHO

POH

O

OHNR3

O

OHHNR3

ONAE

HO

OHO

POH

O

OHNR3

ONAPE-PLD

PLC PLA2

NAE-P

Phosphatase Lyso-PLDGP-NAEPhosphodiesterase

GP-NAELyso-NAPE

ABH4

In the biosynthesis of 2-AG, arachidonic acid containing phosphatidylinositol (PI,

Scheme 1.3) is cleaved at the sn-1 position by phospholipase C (PLC) to provide the

diacylglycerol (DAG).57 This remaining acyl group at the sn-1 position of DAG is cleaved by

diacylglycerol lipase (DGL), which exists in an α and β form, although evidence suggests DGLα

is important in the biosynthesis of 2-AG.58 Another pathway proceeds through Lyso-PI which is

produced through the cleavage of an acyl group of PI by phospholipase A1 (PLA1). 2-AG is then

produced after PI specific cleavage from lyso PI-PLC.57

30

Scheme 1.3 Biosynthetic pathways of 2-AG

AA

O

OP

O

R

O

OH

O

HO

HO

OHOH

OH

PI

PLC

PLA1

AA

OH

O

O

R

DAG

AA

OH

OP

O

OH

O

HO

HO

OHOH

OH

Lyso PI

Lyso PI-PLCDAGL

O

O

OH

OH

2-AG

OH

O

= AA

1.4 Metabolizing Enzymes

The endocannabinoid system is activated by the endocannabinoids biosynthesized on

demand. There are several metabolizing enzymes that are responsible for the degradation of the

endocannabinoids in order to regulate their endogenous levels. These include fatty acid amide

hydrolase (FAAH), monoacylglycerol lipase (MGL), and N-acylethanolamine-hydrolyzing acid

amidase (NAAA).

1.4.1 Fatty Acid Amide Hydrolase - FAAH

Upon metabolism, anandamide is enzymatically hydrolyzed into arachidonic acid and

ethanolamide.59 This metabolizing enzyme was cloned in 1996 by Cravatt, et al., and

distinguished as FAAH, to encompass the fatty acid amides this enzyme was capable of

31

hydrolyzing.60 FAAH was established as the primary metabolizing enzyme of anandamide,

when FAAH-/- knockout mice exhibited a 1500% increase in anandamide levels over wild type.61

FAAH belongs to the amidase family of proteins. Its catalytic activity proceeds through

an unusual mechanism where a non-histidine residue is responsible for activation (histidine most

commonly activates the catalytic serine). Lysine 142 acts as the base in FAAH regulated

metabolism. This activates the serine nucleophile for hydrolytic action.62 The structure of FAAH

consists of a twisted β-sheet with eleven strands encompassed by α-helices. The crystal structure

identified two channels that hold the hydrophobic acyl chain, called the acyl chain binding

channel, and the cytoplasmic-access channel responsible for housing the more polar

ethanolamine.63

With FAAH’s primary function of regulating endocannabinoids, inhibition of this

enzyme should subsequently increase EC levels. This therapeutic potential has seen promise

with regards to neurodegenerative diseases. Endocannabinoids are produced in response to

neurologic trauma of the brain, and an increase in ECs can produce additional therapeutic

relief.64 Inhibition of FAAH has also shown effects toward nicotine reward and dependence.

Besides increasing levels of AEA, inhibition also increases levels of oleoylethanolamide (OEA)

and palmitoylethanolamide (PEA) which can interact with other receptors.65 D’Addario, et al.,

reported that up-regulation of FAAH mRNA is observed in subjects with Alzheimer’s disease

(AD), which results in increased metabolites of AEA, and may lead to increased inflammation

that occurs in AD.66

Phenylmethylsulfonyl fluoride (PMSF, Figure 1.9) was one of the first compounds used

for the inhibition of FAAH. PMSF is a non-selective serine protease inhibitor with an IC50 value

of 290 nM towards FAAH, where pretreatment of this inhibitor has shown to increase AEA

32

levels.67 URB597 is a potent selective irreversible carbamate inhibitor of FAAH with an IC50 of

4.6 nM; while the reversible FAAH inhibitor OL-135 has an IC50 of 2.1 nM.68

SO

OF

PMSF

H2N O

OHN

O

URB597

O

N

O

N

OL-135

Figure 1.9 FAAH inhibitors

1.4.2 Monoacylglycerol Lipase – MGL

MGL was first purified in 1976 from rat adipose tissue and found to be inactivated by

sulfhydryl agents.69 Karlsson, et al., cloned MGL and reported the complete amino acid

sequence containing 302 amino acids. They also identified the catalytic triad of serine 122,

aspartic acid 239, and histidine 269.70 The report of a MGL crystal structure indicates it exists as

a dimer with specific hydrophobic and hydrophilic channels that accommodate 2-AG for

hydrolysis. Cysteine 201 was also identified as a key residue involved in the inhibition of this

metabolizing enzyme.71

Blankman, et al., reported that 85% of all 2-AG metabolism was a result of MGL

activity, with a majority of the remaining hydrolysis mediated by α/β-hydrolase domain-

containing protein-6 and -12.72 Increasing the levels of 2-AG through MGL inhibition produced

behavioral effects in mice similar to those observed with CB1 agonists, such as, analgesia,

hypomotility, and hypothermia.73

One of the more potent and most commonly studied MGL inhibitors is JZL184 (Figure

1.10) with an IC50 value of 8 nM. This inhibition produces cannabinoid behavioral effects such as

analgesia, hypomotility and hypothermia.73

33

O2N

O O

N

OH

O

OO

O

JZL184

Figure 1.10 MGL inhibitor JZL184

1.4.3 N-Acylethanolamine-Hydrolyzing Acid Amidase – NAAA

After the initial identification,74 cloning and characterization,75 NAAA has been

identified as the primary enzyme responsible for PEA hydrolysis, as well as hydrolyzing OEA

and AEA, in many mammalian tissues, organs, and some components of the immune system

(e.g. macrophages).74-77 While NAAA is responsible for the hydrolysis of amides, it is distinctly

different from FAAH.78 These enzymes differ in their primary amino acid sequence and

enzymatic properties. While NAAA is present in the lysosomes, FAAH is primarily distributed

in the cytosolic and luminal sides of intracellular membranes.79 These enzymes appear to have

similar roles, however, their primary substrates are different. NAAA hydrolyzes PEA at a rate

approximately forty times greater than it does AEA,75 while FAAH hydrolyzes PEA eight times

slower than AEA.80 The activity of NAAA is greatest at pH 4-5, whereas FAAH’s peak activity

occurs at pH 8.5-10.81 NAAA is an N-terminal nucleophile hydrolase with a cysteine (Cys126 in

the human enzyme) residue serving as the catalytic nucleophile,75 confirmed by mutagenesis82

and mass spectrometry studies.83

The increase of PEA and OEA through inhibition of NAAA causes increased activatation

of the peroxisome proliferator-activated receptor-α (PPAR-α).84, 85 PEA activation of PPAR-α

produces anti-nociceptive and anti-inflammatory effects.86 While PEA’s activity at the CB1 and

34

CB2 is unclear, it has been hypothesized that PEA participates in an ‘entourage effect’ where

NAEs compete for the active sites of FAAH, and therefore increase the biological activity of

anandamide by impeding its degradation.87 Current reported inhibitors of NAAA are discussed in

Chapter 5.

1.5 Endocannabinoid System Retrograde Signaling

The endocannabinoid system operates under a retrograde signaling pathway.88

Endocannabinoid biosynthesis can be triggered through three mechanisms; Ca2+ elevation

through strong depolarization, Gq-coupled receptor activation, or mild depolarization along with

mild receptor activation.89 Calcium level increase activates the enzymes responsible for

endocannabinoid biosynthesis.90 AEA and 2-AG then migrate after postsynaptic release to

activate presynaptic CB1 receptors.91 This receptor activation then closes Ca2+ channels,

activates K+ channels, and inhibits neurotransmitter release, which stops endocannabinoid

production.89 AEA is then metabolized by FAAH, and 2-AG metabolized by MGL. This

retrograde signaling has implications in areas such as, motor control,92 memory,93 and

neuropathological states.94

1.6 Goal of Thesis Research

The previous review describes important features of the endocannabinoid system that will be

focused on in the following work. The tone of the endocannabinoid is a system can be affected

through a change to receptors, enzymes or biosynthesis/metabolism of endogenous ligands that

work within the system. The compounds prepared here interact with many components in the

endocannabinoid system. Endogenous ligands (AEA and 2-AG) activate both the CB1 and CB2

receptors, while enzymes like FAAH, MGL, and NAAA can directly modify their levels while

also affecting the levels of secondary endocannabinoids.

35

Anandamide based ligands can activate both receptors and is hydrolyzed by FAAH and

NAAA. It is our goal to understand the interactions between this AEA and the CB receptors. It

is our hypothesis that modifying the anandamide head group and altering the tail length of the

ligand combined with a covalent probe can increase the compounds ability to covalent label the

receptor. These interactions are incredibly useful for ligand-assisted protein structure (LAPS)

experiments. This information will help map out the functional selectivity of endocannabinoids

compared to that of other classes of compounds that can activate the CB receptors. We believe

that modifying the head group of the probes can also increase metabolic stability of amide based

ligands in the presence of FAAH, likewise, the same goal is set for analogs of 2-AG.

2-AG the second principle endocannabinoid is difficult to study due to chemical and

metabolic instabilities. Our hypothesis is that through modification of the ligand, we can

produce analogs that will be stable in the presence of metabolizing enzymes so that a

pharmacological profile can be observed.

NAAA can hydrolyze AEA as well as PEA, and it has been observed that affecting the

levels of these endogenous ligands has implications to other endocannabinoids involvement with

FAAH. Our hypothesis is that through the inhibition of NAAA we can increase the therapeutic

effects observed from increased levels of both AEA and PEA. We believe various classes of

compounds (carbamates, carbonates, and isothiocyanates) will behave as potent covalent

inhibitors of the enzymes catalytic cysteine residue.

36

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Endocannabinoid 2-Arachidonoylglycerol Produced by Diacylglycerol Lipase α Mediates Retrograde Suppression of Synaptic Transmission. Neuron 2010, 65, 320-327. 59. Deutsch, D. G.; Chin, S. A. Enzymatic synthesis and degradation of anandamide, a cannabinoid receptor agonist. Biochemical Pharmacology 1993, 46, 791-796. 60. Cravatt, B. F.; Giang, D. K.; Mayfield, S. P.; Boger, D. L.; Lerner, R. A.; Gilula, N. B. Molecular characterization of an enzyme that degrades neuromodulatory fatty-acid amides. Nature 1996, 384, 83-7. 61. Lichtman, A. H.; Shelton, C. C.; Advani, T.; Cravatt, B. F. Mice lacking fatty acid amide hydrolase exhibit a cannabinoid receptor-mediated phenotypic hypoalgesia. Pain 2004, 109, 319-27. 62. Patricelli, M. P.; Cravatt, B. F. Fatty acid amide hydrolase competitively degrades bioactive amides and esters through a nonconventional catalytic mechanism. Biochemistry 1999, 38, 14125-30. 63. Bracey, M. H.; Hanson, M. A.; Masuda, K. R.; Stevens, R. C.; Cravatt, B. F. Structural adaptations in a membrane enzyme that terminates endocannabinoid signaling. Science 2002, 298, 1793-6. 64. Micale, V.; Mazzola, C.; Drago, F. Endocannabinoids and neurodegenerative diseases. Pharmacological Research 2007, 56, 382-92. 65. Muldoon, P. P.; Lichtman, A. H.; Parsons, L. H.; Damaj, M. I. The role of fatty acid amide hydrolase inhibition in nicotine reward and dependence. Life Sciences 2012. 66. D'Addario, C.; Di Francesco, A.; Arosio, B.; Gussago, C.; Dell'osso, B.; Bari, M.; Galimberti, D.; Scarpini, E.; Altamura, A. C.; Mari, D.; Maccarrone, M. Epigenetic regulation of Fatty Acid amide hydrolase in Alzheimer disease. PLoS One 2012, 7, e39186. 67. Wiley, J. L.; Dewey, M. A.; Jefferson, R. G.; Winckler, R. L.; Bridgen, D. T.; Willoughby, K. A.; Martin, B. R. Influence of phenylmethylsulfonyl fluoride on anandamide brain levels and pharmacological effects. Life Sciences 2000, 67, 1573-83. 68. Boger, D. L.; Miyauchi, H.; Du, W.; Hardouin, C.; Fecik, R. A.; Cheng, H.; Hwang, I.; Hedrick, M. P.; Leung, D.; Acevedo, O.; Guimaraes, C. R.; Jorgensen, W. L.; Cravatt, B. F. Discovery of a potent, selective, and efficacious class of reversible alpha-ketoheterocycle inhibitors of fatty acid amide hydrolase effective as analgesics. Journal of Medicinal Chemistry 2005, 48, 1849-56. 69. Tornqvist, H.; Belfrage, P. Purification and some properties of a monoacylglycerol-hydrolyzing enzyme of rat adipose tissue. Journal of Biological Chemistry 1976, 251, 813-9. 70. Karlsson, M.; Contreras, J. A.; Hellman, U.; Tornqvist, H.; Holm, C. cDNA cloning, tissue distribution, and identification of the catalytic triad of monoglyceride lipase. Evolutionary relationship to esterases, lysophospholipases, and haloperoxidases. Journal of Biological Chemistry 1997, 272, 27218-23. 71. Labar, G.; Bauvois, C.; Borel, F.; Ferrer, J. L.; Wouters, J.; Lambert, D. M. Crystal structure of the human monoacylglycerol lipase, a key actor in endocannabinoid signaling. ChemBioChem 2010, 11, 218-27. 72. Blankman, J. L.; Simon, G. M.; Cravatt, B. F. A Comprehensive Profile of Brain Enzymes that Hydrolyze the Endocannabinoid 2-Arachidonoylglycerol. Chemistry & Biology 2007, 14, 1347-1356. 73. Long, J. Z.; Li, W.; Booker, L.; Burston, J. J.; Kinsey, S. G.; Schlosburg, J. E.; Pavon, F. J.; Serrano, A. M.; Selley, D. E.; Parsons, L. H.; Lichtman, A. H.; Cravatt, B. F. Selective

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blockade of 2-arachidonoylglycerol hydrolysis produces cannabinoid behavioral effects. Nat Chem Biol 2009, 5, 37-44. 74. Ueda, N.; Yamanaka, K.; Yamamoto, S. Purification and characterization of an acid amidase selective for N-palmitoylethanolamine, a putative endogenous anti-inflammatory substance. J. Biol. Chem. 2001, 276, 35552-35557. 75. Tsuboi, K.; Sun, Y. X.; Okamoto, Y.; Araki, N.; Tonai, T.; Ueda, N. Molecular characterization of N-acylethanolamine-hydrolyzing acid amidase, a novel member of the choloylglycine hydrolase family with structural and functional similarity to acid ceramidase. Journal of Biological Chemistry 2005, 280, 11082-11092. 76. Sun, Y. X.; Tsuboi, K.; Zhao, L. Y.; Okamoto, Y.; Lambert, D. M.; Ueda, N. Involvement of N-acylethanolamine-hydrolyzing acid amidase in the degradation of anandamide and other N-acylethanolamines in macrophages. Biochimica Et Biophysica Acta 2005, 1736, 211-20. 77. Wang, J.; Zhao, L. Y.; Uyama, T.; Tsuboi, K.; Wu, X. X.; Kakehi, Y.; Ueda, N. Expression and secretion of N-acylethanolamine-hydrolysing acid amidase in human prostate cancer cells. J Biochem 2008, 144, 685-90. 78. Ueda, N.; Yamanaka, K.; Yamamoto, S. Purification and Characterization of an Acid Amidase Selective for N-Palmitoylethanolamine, a Putative Endogenous Anti-inflammatory Substance. Journal of Biological Chemistry 2001, 276, 35552-35557. 79. Ueda, N.; Tsuboi, K.; Uyama, T. N-acylethanolamine metabolism with special reference to N-acylethanolamine-hydrolyzing acid amidase (NAAA). Progress in Lipid Research 2010, 49, 299-315. 80. Wei, B. Q. Q.; Mikkelsen, T. S.; McKinney, M. K.; Lander, E. S.; Cravatt, B. F. A second fatty acid amide hydrolase with variable distribution among placental mammals. Journal of Biological Chemistry 2006, 281, 36569-36578. 81. Ueda, N.; Tsuboi, K.; Uyama, T. N-acylethanolamine metabolism with special reference to N-acylethanolamine-hydrolyzing acid amidase (NAAA). Progress in Lipid Research 2011, 49, 299-315. 82. Wang, J.; Zhao, L. Y.; Uyama, T.; Tsuboi, K.; Tonai, T.; Ueda, N. Amino acid residues crucial in pH regulation and proteolytic activation of N-acylethanolamine-hydrolyzing acid amidase. Biochimica Et Biophysica Acta 2008, 1781, 710-7. 83. West, J. M.; Zvonok, N.; Whitten, K. M.; Wood, J. T.; Makriyannis, A. Mass Spectrometric Characterization of Human N-Acylethanolamine-hydrolyzing Acid Amidase. J Proteome Res 2012, 11, 972-81. 84. Fu, J.; Gaetani, S.; Oveisi, F.; Lo Verme, J.; Serrano, A.; Rodriguez De Fonseca, F.; Rosengarth, A.; Luecke, H.; Di Giacomo, B.; Tarzia, G.; Piomelli, D. Oleylethanolamide regulates feeding and body weight through activation of the nuclear receptor PPAR-alpha. Nature 2003, 425, 90-3. 85. LoVerme, J.; La Rana, G.; Russo, R.; Calignano, A.; Piomelli, D. The search for the palmitoylethanolamide receptor. Life Sciences 2005, 77, 1685-98. 86. Lo Verme, J.; Fu, J.; Astarita, G.; La Rana, G.; Russo, R.; Calignano, A.; Piomelli, D. The Nuclear Receptor Peroxisome Proliferator-Activated Receptor-α Mediates the Anti-Inflammatory Actions of Palmitoylethanolamide. Molecular Pharmacology 2005, 67, 15-19. 87. Lambert, D. M.; Vandevoorde, S.; Jonsson, K.-O.; Fowler, C. J. The Palmitoylethanolamide Family: A New Class of Anti-Inflammatory Agents ? Current Medicinal Chemistry 2002, 9, 663.

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88. Ahn, K.; McKinney, M. K.; Cravatt, B. F. Enzymatic Pathways That Regulate Endocannabinoid Signaling in the Nervous System. Chemical Reviews 2008, 108, 1687-1707. 89. Hashimotodani, Y.; Ohno-Shosaku, T.; Kano, M. Endocannabinoids and Synaptic Function in the CNS. The Neuroscientist 2007, 13, 127-137. 90. Hardie, R. C.; Muallem, S. Lipids in Ca2+ signalling--an introduction. Cell Calcium 2009, 45, 517-20. 91. Wilson, R. I.; Nicoll, R. A. Endogenous cannabinoids mediate retrograde signalling at hippocampal synapses. Nature 2001, 410, 588-92. 92. El Manira, A.; Kyriakatos, A. The role of endocannabinoid signaling in motor control. Physiology (Bethesda) 2010, 25, 230-8. 93. Atsak, P.; Roozendaal, B.; Campolongo, P. Role of the endocannabinoid system in regulating glucocorticoid effects on memory for emotional experiences. Neuroscience 2012, 204, 104-16. 94. Orgado, J. M.; Fernandez-Ruiz, J.; Romero, J. The endocannabinoid system in neuropathological states. Int Rev Psychiatry 2009, 21, 172-80.

43

CHAPTER 2

NOVEL ENDOCANNABINOID PROBES

44

2.1 Introduction

B.R. Baker helped pioneer the idea of covalent ligands in the 1960’s from his study on

enzyme inhibitors, and his pursuits of improving chemotherapy. Baker’s rationale for design was

to understand what chemical features of a ligand were essential to binding so that other chemical

regions of the ligand could be altered to covalently interact with amino acid residues.1 By

incorporating a reactive chemical moiety capable of covalent linkage with amino acid residues,

one could produce ligands that behave as irreversible enzyme inhibitors or as probes to

covalently modify receptor binding sites.

While many functional groups can partake in a myriad of chemical reactions, an effective

strategy for this approach includes reactive functional groups stable in the presence of water and

ones where covalent interaction with specific amino acid residues is selective. These covalent

linkages are most likely to occur with nucleophilic amino acid residues in the active site of an

enzyme; thus electrophilic groups make promising probes.2 Early on, photoaffinity labels were

the most readily used, where photolysis of certain functional groups would cause carbene and

nitrene formation and immediate reactions with nearby amino acid residues would occur.3 One

drawback to this method is the possible promiscuity of carbenes and nitrenes, in which the site of

attachment may be difficult to determine.

Our lab has designed a method utilizing covalent ligands, CB1 and CB2 mutants, along

with LC/MS based methods to characterize the cannabinoid receptors called Ligand-Assisted

Protein Structure (LAPS).4-6 This approach provides insight into the structural and functional

properties of the cannabinoid GPCRs by identifying key amino acid residues in the binding sites

of the receptors. Covalent ligands form an irreversible bond in the binding domain and analysis

45

of the ligand-receptor complex maps out key residues involved in interactions between ligands

and the receptor.

Our laboratory has utilized two types of functional groups in designing covalent

ligands—the azide and isothiocyanate. The azide is a photoaffinity label that when exposed to

irradiation nitrogen gas is lost and a nitrene is formed (Scheme 2.1). This nitrene is then able to

react with nearby amino acid residues in the binding domain to form a ligand-receptor complex.

Although it is observed that azide probes specifically interact with cysteine residues, this may

indicate the a nitrene rearrangement to an imine prior to nucleophilic attack. The isothiocyanate

is an electrophilic group that is unreactive in water, however, it is susceptible to nucleophilic

attack from cysteine to form a covalent bond in the binding domain (Scheme 2.1).

Scheme 2.1: Possible mechanism of nitrene formation and nucleophilic attack of isothiocyanate

R NN

NR N

NN

hv

-N2

R N R NC

SNu

H

R NH

Nu S

2.2 Current Cannabinoid Probes

The first cannabinoid probe to covalently bind to the CB1 receptor was (-)-5`-azido-Δ8-

tetrahydrocannabinol (5`-azido-THC, 1, Figure 2.1). The Ki for 1 to the CB1 receptor was 19

nM, which was a two-fold increase in affinity compared to the parent (-)-Δ8-THC (Ki of 35 nM).

The covalent binding of 1 was 12% and 31% for 50nM and 500nM equilibrated concentrations,

respectively, after 5 minutes of UV exposure.7

46

O

OH

N3 O

OH

1 2

OH

NCS

O

OH

NCS

3

O

Figure 2.1 THC based covalent probes

(-)-11-hydroxy-7`-isothiocyanato-1`,1`-dimethylheptyl-Δ8-THC (2, Figure 2.1) was

developed as an electrophilic CB1 covalent probe. The IC50 of 2 was measured to be 1.6 nM. At

0.5 nM 10% labeling was observed after 5 min incubation and 50% after 30 min incubation.

Labeling increased to 50% and 70% labeling for 5 and 60 min incubation respectively using a 2

nM concentration of 2.8 (–)-7′-isothiocyanato-Δ8-THC (3, Figure 2.1) was synthesized in 1995

showing similar binding affinity and covalent binding data to 2.9 At that time, the covalent

binding of the isothiocyanate probes lead to the conclusion that a thiol, amine, or imidazole

amino acid was in the vicinity of the active site. In 2005, it was discovered that C6.47 (meaning

cysteine on helix 6, which is the 47th amino acid residue of the helix, it is also the 355th amino

acid residue overall) was the site of covalent attachment of 2 (AM841) to the CB1 receptor.

Treatment of 2 with alanine, serine, and leucine mutants of C6.47 (355) showed no change in

binding of [3H]CP55940 and [3H]WIN55212-2 to the CB1 receptor, whereas, treatment of 2 with

the WT CB1 receptor eliminated binding of [3H]CP55940 and [3H]WIN55212-2.10

47

Photoactivatable anandamide analog 4 (Figure 2.2) was designed as a potential probe for

the identification of non-CB1 and non-CB2 receptors. Compound 4 has a Ki of 570 nM for CB1,

and 220 nM for CB2. The reduced affinity to the CB receptors was thought to allow this probe to

identify other non-CB receptors for anandamide.11 Covalent probe 5 (Figure 2.2) was an

improvement on 3 in which an iodo-group was incorporated for potential radiolabeling. This

compound was a successful probe where at 25 nM 50% of 5 was covalently bound to the

receptor.

O

NH

O O

N3

I

4

O

O

OH

NCSI

5

OH

Figure 2.2 Other endocannabinoid probes

Martin-Couce, et al., developed a series of endocannabinoid probes for labeling of the

CB receptors. These ligands will had one of three tags; biotin, benzophenone, or a terminal

alkyne attached to a backbone analogous of AEA, 2-AG, and 2-AGE (Figure 2.3). Ligands 6-10

had poor affinity for the CB1 and CB2 receptors with Ki values >2000 nM. The 2-AG and 2-

AGE based ligands did, however, show some affinity to the CB receptors with compound 11

exhibiting Ki values of >5000 nM (CB1) and 379 nM (CB2); 12: 221 nM (CB1) and 450 nM

(CB2); and 13: 84.7 nM (CB1) and 84.9 nM (CB2).12

48

O

NH

OTag1

O

NH

OH

OTag1

Tag1 =

ONH

O

5 3

S

HN NHHH

O

Tag2 =

O3

Tag3 =

O

HN

O

3

O

O

NH

OO

NH

OO

NH

OTag1 Tag2 Tag3

O3

OH

OTag1 O

OH

OTag1 O

OH

OTag2

O

6 7

8 9 10

11 12 13

Figure 2.3 Tagged endocannabinoid probes

The first two high affinity covalent anandamide-based probes for the CB1 receptor were

previously synthesized by our lab in 2005. Ligands 14 and 15 (Figure 2.4) were synthesized

with either the azide or isothiocyanate covalent probe at the ω-6 position and a cyclopropylamide

at the head position. These ligands showed a very high affinity towards the CB1 and the CB2

receptor; with 14 exhibiting Ki values of 0.9 nM (CB1) and 58 nM (CB2), and 15 exhibiting Ki

values of 1.3 nM (CB1) and 49 nM (CB2). After equilibrating 14 with the membranes at a

concentration 10 times its Ki, the membranes were exposed to ultraviolet light (254 nm) and 14

49

was irreversibly labeled at 42%. Electrophilic probe 15 equilibrated with the receptor at a

concentration 10 times its Ki, irreversible labeled 32% of the receptors.13

O

NH

N3

O

NH

SCN14 15

Figure 2.4 Arachidonoylcyclopropylamine covalent probes

2.3 Design of Bifunctional Endocannabinoid Covalent Probes

Endocannabinoid covalent probes had previously exhibited high affinity towards the CB

receptors and both photoaffinity and electrophilic probes were shown to irreversibly bind to the

rCB1 receptor. The next step was to synthesize bifunctional probes with increased covalent

binding to rCB1 (rat), mCB2 (mouse), and hCB2 (human). Finding new compounds with

enhanced covalent binding combined with covalent studies on CB receptor mutants can reveal

important ligand-protein interaction in the binding domain for this class of compounds. This

information can enhance the design and synthesis of more potent ligands.

The design of our probes was based on key functional groups from prior

endocannabinoid analogs. The azide probe of 14 and the isothiocyanate probe of 15 were to be

used at the tail for irreversible linkage, while the head group would be a terminal alkyne as seen

with 16 (Scheme 2.2). 16 has high affinity for both CB receptors with Ki values of 10.8 nM

(rCB1 with PMSF), 4900 nM (rCB1), and 290 nM (mCB2).14 The terminal alkyne head-group

has shown an unexpected improvement to higher binding affinity at the CB2 receptor while also

introducing a second functionality to the molecule. The alkyne can be used for subsequent

“click” chemistry reactions that may be useful in the purification of a receptor-ligand complex.

50

We hypothesized that incorporating these functional groups while altering the arachidonic chain-

length would result in improved covalent labeling of the cannabinoid receptors.

Scheme 2.2 Design of bifunctional azide and isothiocyanate covalent probes

O

NH

16

O

NH

X14 X = N315 X = NCS

nX

O

NH

n = 2-5

17-20 X = N321-24 X = NCS

2.4 Linear synthetic strategy

The synthetic route originally employed for 14 and 15 proceeded through a linear

synthesis, utilizing a series of 1,4-skipped alkyne couplings from a terminal alkyne and

propargyl halide15 to eventually produce 20-hydroxy methyl tetraynoate (Scheme 2.3).

Drawbacks of this approach include the instability of the sequential 1,4-skipped alkyne

intermediates. If a synthesis could be developed that may limit the amount of intermediate 1,4-

skipped alkynes, and number of steps, the overall yield can be improved.

Scheme 2.3 Linear synthesis of 20-hydroxy methyl tetraynoate

HOO

OHO

ClO

O

BrO

OOH

O

OHO

O

OBr

OH O

OHO

CuI, NaI, K2CO3,DMF, rt

CBr4, PPh3, CH2Cl2, 0 °C




51

2.5 Development of a Convergent Synthesis for Covalent Probes

An efficient way to improve linear syntheses is to develop a convergent strategy. A

scheme was devised to synthesize two diyne intermediates that would resemble the two halves

(head and tail) of the probe based on a method reported in the synthesis of 4.11 In doing so, the

triyne intermediate, whose chemical instability causes a loss in the final yield, would be avoided.

The modified synthesis of the tetraynoate (38-41, Scheme 2.4) started with the coupling of

trimethylsilyl (TMS) protected propargyl bromide 25 to alkynols 26-29 in the presence of CuI,

NaI, and K2CO3 to yield TMS protected diynes 30-33 in 54-73% yields. As in previous

methods, 30 was deprotected with TBAF to yield deca-6,9-diyn-1-ol (Scheme 2.5). This method

proved to be successful on a scale <2.25 mmol where the yield was consistently >90%.

However, in a scale up of the synthesis, deprotection of 30 produced large amounts of an

unwanted allene byproduct. This drastically reduced the yield for the total synthesis, as a major

loss of compound occurred here. Montel et al. reported a 1,4-skipped alkyne synthesis from

TMS-alkynes and propargyl halides with CuI, and a fluorine source that allowed the deprotection

and coupling to occur in a one-pot synthesis.16 Thus, after CuI coupling of 34 and 35 with

subsequent Appel17 reaction to yield bromide 36 in 63% over the two steps, the TMS protected

diynes 30-33 were coupled to 36 by introducing 1.0M TBAF in THF (CsF as the fluorine source

reacted much slower and resulted in lower yields) to the CuI coupling conditions to yield

tetraynoates 38-41 in 34-63% yields.

52

Scheme 2.4 Convergent synthesis of tetraeneoates

O

OHOCl

OHTMS

Brn

n = 2-5

OH

n

TMS

HO

O

OBr

O

OHO

OH

O

O

n = 2-5n

O

O

nn = 2-5 n = 2-5


25 26-29 30-33

34 35 36

3730-33

38-41 42-45

54-73%


74%


85%

CuI, NaI, K2CO3, CsF, DMF, rt

34-63%

Ni(OAc)2, NaBH4, NH2(CH2)2NH2

33-57%

Scheme 2.5 Deprotection of 10-(trimethylsilyl)deca-6,9-diyn-1-ol

TMSOH

HOH

.5eq TBAF, THF-40 to 0oC A

B

C D

53

Partial hydrogenation of tetraynoates (38-41) is a crucial step as the synthesis proceeds

from the most unstable intermediates to the most stable intermediates, tetraenoates 42-45.

Initially, the partial hydrogenation was completed through Lindlar’s catalyst and quinoline,18

however, on a compound containing four alkynes, mixed saturation products were observed.

Many different reaction parameters were attempted to optimize this reaction, as it is simple to

carry out. However, after adjusting reaction temperature, equivalents of quinoline, and reaction

concentrations, no conditions proved satisfactory and it was apparent that the Lindlar catalyst

was more suited for partial hydrogenation of compounds containing one or two alkynes. A

slightly more involved partial hydrogenation reaction was then used. A P-2 Ni catalyst was

formed in situ from nickel(II) acetate tetrahydrate, sodium borohydride, and ethylenediamine

which was able to hydrogenate 38-41 to the all cis-tetraenoates 42-45.19

To reach the final products, the tail end hydroxyl was converted to an azide with

diphenylphosphoryl azide (DPPA) and 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) to produce

46-49 in ~55% yields (Scheme 2.6). The methyl esters were then hydrolyzed to the acids 50-53

in 98% yield. 50-53 then went through carbodiimide induced amide coupling with propargyl

amine to produce the azide probes 17-20 in ~80% yield. Subsequent treatment with PPh3 and

CS2 converted the azides to the isothiocyanates to provide probes 21-24.

Scheme 2.6 Functionalizing the arachidonoate tail

nN3

O

O

n = 2-5nN3

O

OH

n = 2-546-49 50-53

nN3

O

NH

n = 2-517-20nNCS

O

NH

n = 2-521-24

DPPA, DBU, DMF, 120 °C

~55%

LiOH, THF, rt

~80%

propargylamine, EDCI, DMAP, DCM, 0 °C

~80%

CS2, PPh3, THF, rt

~60%

42-45

54

2.6 Binding Affinity and Covalent Binding Data of Covalent Probes

It has been established that anandamide analogs with shortened tail length can activate

the CB receptors. Yao, et al., synthesized methanandamide (AM356) analogs varying the tail

length of the compound while introducing a terminal aryl group. The binding affinity of those

compounds decreased when the tail was shortened to two carbons beyond the C14-C15 olefin.20

The same trend was observed with ligands 17 and 21. Where n=2, the binding affinity

was ~100 nM for the rCB1 receptor and >300 nM and >130 nM for the mCB2 and hCB2

receptors respectively (Table 2.1). For the ligands where n=3,4, or 5, the binding affinity is

retained, with ligands 19 and 20 having affinities in the low nM range for rCB1 with PMSF.

Another very interesting result was the binding affinity of 19 to the hCB2 receptor; at 15 nM it is

has one of the lowest Ki values for a lipid based compound toward the CB2 receptor.

aData obtained from one experiment run in triplicate. bData obtained from at least two separate experiments and no more than five all run in triplicate.

Table 2.1 Binding affinity (Ki) for ligands 17-24 to the cannabinoid receptors

nX

O

NH

Ligand AM n X rCB1 (nM) rCB1 w/

PMSF (nM) mCB2 (nM) hCB2 (nM)

17 9025a 2 N3 881.7 110.1 443.8 132.1

18 9032a 3 N3 121 21 38±1.3 36±2.6

19 9014b 4 N3 350±32 7.4±1.2 34±4.0 22±2.7

20 9002a 5 N3 >1000 6.9 43 55

21 9029b 2 NCS 1600±160 85±9.3 300±33 190±23

22 9039b 3 NCS 280±57 17±2.7 51±15 28±4

23 9017b 4 NCS 560±49 17±0.6 44±5.1 43±8.6

24 9004a 5 NCS >1000 12 25 25

55

The increased potency for these compounds towards the CB2 receptor was not originally

expected, as LAPS studies on the CB2 receptor using endocannabinoid templates would bring

new insight to their interactions with that receptor. Ideally, for covalent studies one would

prefer the Ki to be ≤30 nM. As our covalent assays use a standard concentration of 10Ki (10

times the concentration of the Ki), the greater the affinity the more likely we can avoid solubility

issues. Using these criteria, ligands 17 and 21 were considered not suitable for covalent binding

studies for either receptor, while all the other ligands had acceptable binding affinities.

Azide probes 18-20 did not show any promising covalent activity towards CB2 receptors,

with only 19 successfully labeling the mCB2 receptor (20%, Table 2.2). Isothiocyanates 22-24

showed covalent binding greater than that of previous probes 14 and 15 (62-78% vs. 32-40%

respectively). The covalent binding to the CB2 receptors was greatly improved with 23, where

the ligand labeled 53% of the mCB2 receptor and 38% of hCB2; thus, making it a good candidate

for LAPS studies with the CB2 receptor. CB1 cAMP assay of 23 indicated that compound

behaved as an agonist with an EC50 of 85 nM.

aAll covalent binding data was obtained from one experiment run in triplicate.

Table 2.2 Percentage of occupied receptors for 18-20 and 22-24 in covalent binding assaya

nX

O

NH

Ligand AM n X rCB1 mCB1 hCB2

18 9032 3 N3 - 0% 0%

19 9014 4 N3 - 20% 4%

20 9002 5 N3 - 0% 0%

22 9039 3 NCS 78% 32% 27%

23 9017 4 NCS 62% 53% 38% 24 9004 5 NCS 71% 29% 24%

56

Even with optimal receptor binding it is difficult to predict which ligands will produce

increased receptor labeling. The azide and isothiocyanate groups are linear, and placing them at

the tail of a compound increases the overall length of the tail and reduces the degrees of freedom

that may be required for adequate receptor binding (Scheme 2.7).

To circumvent this possibility we investigated shortening the arachidonic tail where the

probe would be attached. We hypothesized that reducing the tail length, while incorporating the

probe, would produce similar affinity to that of the ω-6 ligands. It was thought the shorter chain

would then place the probe in a more advantageous region for covalent linkage. However, this

trend was not observed as ligands 22-24 exhibited similar covalent binding percentages; and the

only difference in covalent binding was observed between the use of azide and isothiocyanate

probes themselves. This may be due to the mechanism of azide irradiation and rearrangement, or

the UV irradiations affect on the receptor itself.

Scheme 2.7 Crude visualization of linear probes degree of freedom

14 15 14 15N

CS

NNC

SCS

2.7 Head-group Optimization

One drawback to anandamide based probes is their susceptibility to hydrolysis by FAAH.

CB1 assays are run with the FAAH inhibitor PMSF to prevent amide hydrolysis.

Methanandamide (AM356) was the first synthetic anandamide analog that was metabolically

stable in the presence of FAAH.21 This metabolic stability occurs from the incorporation of a

chiral methyl to the ethanolamine head group--alpha to the amide nitrogen (Scheme 2.8). In

57

order to obtain the metabolic stability desired towards FAAH we decided to introduce a chiral

methyl onto the propargyl amide head group.

Scheme 2.8 Anandamide and AM356 CB1 binding

O

NH

OHO

NH

OH

rCB1: >1000nMrCB1 w/ PMSF: 78nM

rCB1: 28nMrCB1 w/ PMSF: 20nM

Anandamide AM356

The synthesis of the chiral head group started from the D and L enantiomers of N-BOC-

(tert-butyloxycarbonyl) alaninol (54 and 55, Scheme 2.9). The alcohol was oxidized (Swern22

conditions) to yield 56 and 57, which were immediately reacted with the Ohira-Bestmann23

reagent to produce terminal alkynes 58 and 59 with 30-56% yields over the two steps. The BOC

protecting group was removed with TFA (triflouroacetic acid) to produce salts 60 and 61 in good

yields.

Scheme 2.9 Synthesis of chiral head group

NH

HOO

O NH

OO

O(COCl)2, DMSO, Et3N-78°C DCM

OP

OO

O

N2

MeOH, K2CO3 16h, 0°

NH

O

O

30-56% from alcohol

TFA, DCM NH3 O

O

CF3~90%

54 = R55 = S

58 = R59 = S

56 = R57 = S

60 = R61 = S

58

Salts 60 and 61 were then coupled to acid 52 with 1-ethyl-3-(3-

dimethylaminopropyl)carbodiimide (EDCI) and 4-dimethylaminopyridine (DMAP) in

quantitative yield to give the R and S amides, 62 and 63. The azides were easily converted to

isothiocyanates 64 and 65 with previously described conditions (Scheme 2.10).

Competition binding assays indicated that 62-65 were stable in the presence of FAAH.

This was determined by the examining the similarities in binding affinity between the rCB1

assays with PMSF and without. It was surprising to see that the chirality of the methyl group did

not exhibit a significant difference on binding affinity to the CB1 receptor; whereas (R)-

methanandamide had an eight fold greater binding affinity than that of (S)-methanandamide.21

The R-isomers however, did have a higher affinity for the hCB2 receptors, while all ligands saw a

decline in mCB2 affinity compared to other covalent ligands (Table 2.3).

Scheme 2.10 Metabolically stable covalent probes

O

OHN3n n = 4

EDCI, DMAPDCM, 0°C

O

NHN3

n n = 4

100%

PPh3, CS2THF, 48hrs

45-52%

O

NHNCS

n n = 462 = R63 = S

64 = R65 = S

NH3 O

O

CF3

60 = R61 = S

52

The metabolically unstable isothiocyanate probes were better candidates for covalent

binding studies by occupying a much larger percentage of binding sites compared to their azide

counterparts. The opposite was observed for the metabolically stable chiral probes. The

isothiocyanate probes showed no covalent labeling at any of the receptors, while (S)-azide probe

63 occupied 77% of available CB1 receptors in the covalent binding assay. The (R)-azide, 62,

59

only yielded 8% covalent binding percentage towards CB1, however, 62 was the best candidate

for hCB2 covalent binding as it occupied 46% of available receptors (Table 2.4). Compound 62

was observed to behave as an agonist at the CB1 receptor with an EC50 of 11 nM, while observed

to behave as an inverse agonist at the hCB2 receptor.

Table 2.3 Binding affinity (Ki) for ligands 62-65 to the cannabinoid receptorsa

O

NH

X

Ligand AM X Chirality rCB1 (nM)

rCB1 w/ PMSF (nM)

mCB2 (nM)

hCB2 (nM)

62 9069 N3 R 5.9±1.8 11±0.4 92±17 37±10

63 9073 N3 S 28±6.8 16±1.1 250±21 96±14

64 9070 NCS R 18±4.4 15±4.4 97±2.1 63±20

65 9074 NCS S 35±7.5 29±7.3 164±25 140±21 aAll experiments done in triplicate in three separate experiments.

Table 2.4 Percentage of occupied receptors by 62-65 in covalent binding assaysa

O

NH

X

Ligand AM X Chirality rCB1 hCB2 62 9069 N3 R 7.6% 46%

63 9073 N3 S 77% -

64 9070 NCS R 0% 0%

65 9074 NCS S 0% - aAll experiments completed once in triplicate.

60

2.8 Ligand-Assisted Protein Structure (LAPS)

LAPS analysis combines the use of covalent probes, mass spectrometry, site-directed

mutagenesis, and computer modeling to determine the binding locations of different classes of

ligands to the CB GPCRs. This technique is used to understand structural and functional

information regarding the interaction of ligands and target proteins. Information obtained can be

used in the development of novel therapeutics.

Covalent analogs to THC have been used to determine within which transmembrane helix

(TMH) these types of ligands interact.10 Endocannabinoids are important signaling compounds,

information related to their interaction with the CB receptors is important in understanding their

function. Compound 15 was one of the first covalent probes analogous to the endocannabinoid

anandamide that showed adequate covalent binding percentages to the CB1 receptor to be used

for LAPS studies.13

To determine where in the GPCR ligands interact, a series of cysteine to serine mutants

of the CB receptors was developed. The CB1 receptor contains five cysteines located within the

transmembrane domains (Figure 2.5) at C1.55(139), C4.47(238), C6.47(355), C7.38(382), and

C7.42(386).

61

Figure 2.5 CB1 receptor with transmembrane cysteines highlighted

With the covalent warhead (azide or isothiocyanate) susceptible to attack only from a

nucleophilic cysteine, it is possible to determine which specific cysteine interacts with the

covalent probe. By sequentially mutating each cysteine to serine and performing a covalent

assay, we can determine the specific cysteine where interaction occurs by looking for a loss of

covalent labeling.

This method was used on rCB1 with 15, and it was observed that the covalent binding

interaction was lost when C6.47(355) was mutated to serine.24 The covalent probe occupied

some percentage of the CB1 receptors when a covalent assay was performed with the wild type

receptor and cysteine to serine mutants at the other helices (C1.55, C4.47, C7.38, and C7.42).

2.8.1 LAPS Studies Utilizing AM9017

LAPS studies of 15 indicated an interaction with the CB1 receptor at TMH 6, however,

this compound was unable to covalently bind with the CB2 receptor. Endocannabinoids (ECs)

62

have a greater affinity for CB1 compared to CB2, thus understanding interactions of ECs using

LAPS can be difficult with the CB2 receptor.

While the original strategy for incorporating a propargyl amine head-group into the EC

probe was for the potential use of ‘click’ chemistry (selective and efficient functional group

bonding) after the ligand covalently labeled to the receptor, we observed an unexpected

improvement of ligand affinity towards the CB2 receptors. With binding affinities in the lower

nM range, we would be able to perform covalent assays at a 10x Ki standard concentration

without solubility issues. The compound with the best covalent activity for both mCB2 and hCB2

was AM9017 (23, Table 2.2), and it would be the best candidate for LAPS studies on mutant

CB2 cell lines.

C7.38(284)SC7.42(288)S

C6.47(257)S

C1.39(40)S

C2.59(89)S

Site-Directed MutagenesishCB2 Library of Cysteine Mutants

http://www.wdv.com/CellWorld/Receptors/

Figure 2.6 Human CB2 receptor with transmembrane cysteines highlighted

63

CB2 also contains five cysteines located in the transmembrane helices. These residues

are located at C1.39(40), C2.59(89), C6.47(257), C7.38(284), and C7.42(288) (Figure 2.6).

AM9017 (23) was tested in covalent assays on the wild type CB2 receptor for mouse and human.

The Bmax of [3H]-CP55,490 with the mCB2 receptor treated with AM9017 was 7100 (units for

Bmax are ρmol mg-1) compared to a Bmax of 15000 for the untreated mCB2 receptor; indicating

AM9017 occupied 53% of the mCB2 receptors. With the hCB2 receptor treated with AM9017,

[3H]-CP55,490 had a Bmax of 800, and a Bmax of 500 for the neat hCB2 receptor. This indicates

that AM9017 occupied 38% of the hCB2 receptors (Figure 2.7).

Covalent binding assays were performed on mutant mouse and human CB2 cell lines.

The first mutant assay to be performed was cysteine to serine on TMH6 (C6.47(257)S). The

Bmax of [3H]-CP55,490 on the neat receptor compared to receptor treated with AM9017 was

similar on both the mouse (7600 for neat compared to 8100 for treated mCB2) and human (6500

for neat compared to 6900 for treated hCB2) CB2 receptors (Figure 2.8). The loss of covalent

labeling of AM9017 when the cysteine on TMH 6 was mutated to a serine indicates this was the

site of interaction between AM9017 and the CB2 receptor.

64

(A)

(B)

Figure 2.7: Competition binding of [3H]-CP55,490 with the (A) mouse and (B) human CB2 wild type receptors. Covalent binding is determined by the difference in competition binding of [3H]-CP55,490 with the neat receptor and the receptor pretreated with AM9017.

65

Figure 2.8: Competition binding of [3H]-CP55,490 with the (A) mouse and (B) human CB2 receptors where C6.47(257) was mutated to S6.47(257) on TMH6. Covalent binding is determined by the difference in competition binding of [3H]-CP55,490 with the neat mutant receptor and the mutant receptor pretreated with AM9017.

66

2.9 Significance of Transmembrane Helix 6

Studies of the β2-adrenergic25 and bovine rhodopsin26 GPCR indicate that the inactive

form of this class of GPCRs contains a salt bridge between TMH3 and TMH6. Upon GPCR

activation the salt bridge is broken due to the increase in distance between involved residues.

TMH6 appears to be significant for the activation of class-A GPCRs. This stems from a CWxP

motif which is conserved throughout this class of GPCRs.27, 28 This structural conformation has

been confirmed to exist in the hCB2 receptor through NMR spectroscopy.29 The tryptophan

(W258) in the conserved kinked region of TMH6 has been identified through NMR to hydrogen-

bond to a carbonyl from a nearby leucine residue (L255). H-bonding then positions the cysteine

(C257) in a favorable position for ligand interaction.29

2.10 Docking of AM9017 into hCB2 Models

Crystal structures have been produced of the rhodopsin30 and the β231 and A2A

32

adrenergic receptors in both their active and inactive states. These crystal structures suggested

that a change in state caused a shift in the cytoplasmic ends of TMH5 and TMH6. We have

modeled AM9017 in the hCB2 receptor based on these GPCR crystal structures.

AM9017 was first non-covalently docked to hCB2 receptor, followed by covalent

attachment of isothiocyanate to cysteine in TMH6. AM9017-hCB2 complex was inserted into a

palmitoyloleoylphosphatidylcholine (POPC) membrane, and the entire system was then

equilibrated with production dynamics of 1200 ps (Figures 2.9 and 2.10). The AM9017-hCB2

complex was then overlayed on the inactive hCB2 receptor (Figure 2.11).

67

Figure 2.9 AM9017-hCB2 covalent complex equilibrated in a POPC membrane

68

Figure 2.10 Zoom-in of AM9017 covalent attachment to the hCB2 receptor with removal of POPC membrane. AM9017 is in stick representation with magenta carbons, blue nitrogen, yellow sulfur, and cysteine residue in orange. hCB2 is represented with green carbons

69

Figure 2.11 Comparison of inactive hCB2 (orange) with active hCB2 receptor covalently bound to AM9017 (green)

70

2.11 Conclusions

To benefit from LAPS studies, a convergent synthesis was developed and optimized to

produce bifunctional anandamide-based covalent probes. This convergent synthetic route

overcame stability issues of skipped-alkyne intermediates observed in the linear synthesis, as

well as overcome the inconsistent partial hydrogenation from using Lindlar’s catalyst. By

modifying the number of carbons in the ligand tail we observed that 3, 4, and 5 carbon tail

lengths were needed for binding affinities high enough for adequate covalent binding using a

standard 10x Ki ligand concentration. These ligands incorporated a propargyl amine head group

which improved CB1 affinity and unexpectedly the CB2 receptor binding as well. AM9017 was

identified as a candidate for LAPS studies of anandamide-based analogs on the hCB2 receptor,

where endocannabinoid covalent binding data had previously been elusive. AM9017 occupied

62, 53, and 38% of the rCB1, mCB2, and hCB2 respected binding sites. Preliminary LAPS

studies on hCB2 mutants, where C6.47 is mutated to serine, showed a loss of covalent labeling

with AM9017. This implied that TMH6 was the site of interaction between covalent probe

AM9017 and the hCB2 receptor. This interaction is believed to occur at a conserved CWxP

motif in TMH 6, which is also observed with anandamide-based covalent probes at the rCB1

receptor. When the ligand-protein covalent interaction was modeled based on crystal structures

of other GPCRs, a movement of TMH6 towards TMH5 was observed.

One of the disadvantages of the bifunctional covalent probes was their susceptibility to

enzymatic hydrolysis by FAAH. To reduce the susceptability, a propargyl amide head group

was designed with a chiral methyl group α to the amide nitrogen. This methyl group was

successful in stabilizing the ligand in the presence of FAAH, as indicated by binding assays in

the presence and absence of the FAAH inhibitor, PMSF. In contrast to the results from previousl

71

anandamide-based compounds that introduced chiral methyl groups for this purpose, the chirality

of the methyl group in these compounds did not have a significant impact on receptor binding.

In summary, new synthetic routes were developed to expedite the preparation of novel

endocannabinoid based ligands, with an emphasis on ligands ability for covalent attachment to

cysteine residues. These advances are crucial for a better understanding of ligand-protein

interactions within the cannabinoid system. This information will be beneficial for the design

and development of future therapeutics targeted for the cannabinoid receptors.

72

2.12 Experimental

General. All starting materials were obtained from commercial suppliers and were used without

further purification. 1H NMR and 13C NMR spectra were obtained on either a Varian 400 or 500

MHz spectrometer with CDCl3 as the solvent and TMS as the internal standard. Signal

multiplicities are labeled as: s for singlet, d for doublet, t for triplet, q for quartet, quin for

quintet, m for multiplet, and any appropriate combinations. All chemical shifts are in ppm. IR

spectra were obtained on a Perkin-Elmer Spectrum One FT-IR spectrometer. All HRMS data

were obtained on a Micromass 70-VSE III mass spectrometer and performed by the School of

Chemical Science, University of Illinois at Urbana-Champaign, Urbana, IL. All compounds were

purified on a Biotage Isolera One using prepacked Luknova flash columns with 40-60μM, 60Å

silica gel. Cannabinoid receptor binding assays and covalent binding assays were conducted

following the procedures outlined by Li et al.; except covalent binding assays were performed

with a concentration of 10Ki of each ligand.13

O

NHN3

(5Z,8Z,11Z,14Z)-17-azido-N-(prop-2-ynyl)heptadeca-5,8,11,14-tetraenamide, 17, AM9025.

EDCI (190 mg, 0.99 mmol), DMAP (80 mg, 0.66mmol) and propargyl amine (36 μL, 0.66

mmol) were added to a solution of 50 (100 mg, 0.33 mmol) in anhydrous CH2Cl2 (5 mL) at 0oC.

The reaction was allowed to stir under argon for 3 h. Upon completion; the reaction was diluted

with CH2Cl2 and water. The organic layer was separated, concentrated, and chromatographed on

silica to yield 17, AM9025, (.102g, 91%) as a yellow oil. Rf = 0.42 (35% ethyl acetate /

hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.90 (m, 1 H) 5.47 - 5.56 (m, 1 H)

5.29 - 5.46 (m, 7 H) 4.02 (dd, J=5.4, 2.4 Hz, 2 H) 3.28 (t, J=7.1 Hz, 2 H) 2.72 - 2.88 (m, 6 H)

73

2.33 - 2.41 (m, 2 H) 2.16 - 2.23 (m, 3 H) 2.10 (q, J=6.8 Hz, 2 H) 1.71 (quin, J=7.5 Hz, 2 H). 13C

NMR (126 MHz, CHLOROFORM-d) δ ppm 172.7, 131.2, 129.3, 129.0, 128.6, 128.5, 128.2,

128.0, 125.6, 79.9, 71.7, 51.2, 35.9, 31.8, 29.3, 27.3, 26.8, 26.0, 25.9, 25.5. IR (neat) cm-1 3296,

2093, 1649, 1533.

O

NH

N3

(5Z,8Z,11Z,14Z)-18-azido-N-(prop-2-ynyl)octadeca-5,8,11,14-tetraenamide, 18, AM9032.

Synthesized following the procedure for 17 from 51 (84 mg, 0.26 mmol) to yield 18, AM9032,

(65 mg, 71%) as a yellow oil. Rf = 0.63 (35% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 5.80 (br. s., 1 H) 5.21 - 5.50 (m, 8 H) 4.05 (dd, J=4.9, 2.4 Hz, 2 H)

3.22 - 3.32 (m, 2 H) 2.73 - 2.86 (m, 6 H) 2.22 - 2.24 (m, 1 H) 2.17 - 2.22 (m, 2 H) 2.04 - 2.15 (m,

4 H) 1.70 - 1.77 (m, 2 H) 1.56 - 1.65 (m, 2 H). 13C NMR (126 MHz, CHLOROFORM-d) δ

172.7, 130.4, 130.1, 129.3, 129.2, 129.1, 128.7, 128.2, 128.1, 79.9, 71.8, 51.7, 35.9, 29.7, 29.4,

29.0, 27.3, 26.9, 25.9, 25.5, 24.5. IR (neat) cm-1 3296, 2094, 1646, 1535.

O

NH

N3

(5Z,8Z,11Z,14Z)-19-azido-N-(prop-2-ynyl)nonadeca-5,8,11,14-tetraenamide, 19, AM9014.

Synthesized following the procedure for 17 from 52 (122 mg, 1.0 mmol) to yield 19, AM9014


CHLOROFORM-d) δ ppm 6.06 (br. s., 1 H) 5.29 - 5.43 (m, 8 H) 4.01 (dd, J=4.9, 2.4 Hz, 2 H)

3.25 (t, J=6.8 Hz, 2 H) 2.71 - 2.86 (m, 6 H) 2.15 - 2.22 (m, 3 H) 2.01 - 2.12 (m, 4 H) 1.70 (quin,

J=7.5 Hz, 2 H) 1.59 (quin, J=6.8 Hz, 2 H) 1.43 (quin, J=7.6 Hz, 2 H). 13C NMR (126 MHz,

74

CHLOROFORM-d) δ ppm 172.8, 129.6, 129.3, 129.0, 128.6, 128.5, 128.4, 128.3, 128.2, 79.9,

71.6, 51.5, 35.8, 29.3, 28.6, 27.3, 27.0, 26.9, 26.8, 25.8 (2 C), 25.56. IR (neat) cm-1 3299, 2090,

1647, 1536.

O

NH

N3

(5Z,8Z,11Z,14Z)-20-azido-N-(prop-2-ynyl)icosa-5,8,11,14-tetraenamide, 20, AM9002.


29 mg, 81%) as a yellow oil. 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.32 - 5.44 (m, 8

H) 4.05 (dd, J=5.4, 2.44 Hz, 2 H) 3.27 (t, J=6.8 Hz, 2 H) 2.75 - 2.88 (m, 6 H) 2.23 (t, J=2.4 Hz, 1

H) 2.19 (t, J=7.45 Hz, 2 H) 2.05 - 2.16 (m, 4 H) 1.73 (quin, J=7.3 Hz, 2 H) 1.60 (quin, J=6.45

Hz, 2 H) 1.36 - 1.44 (m, 4 H); IR (neat) cm-1 3297, 2094, 1647, 1533. 13C NMR (126 MHz,


71.5, 51.9, 35.8, 29.3, 28.6, 27.3, 27.0, 26.9, 26.8, 25.9, 25.7 (2 C), 25.5. HRMS for C23H35N4O

(MH+) 383.2803. Calcd. 383.2811.

O

NHNCS

(5Z,8Z,11Z,14Z)-17-isothiocyanato-N-(prop-2-ynyl)heptadeca-5,8,11,14-tetraenamide, 21,

AM9029. Triphenylphosphine (92 mg, 0.35 mmol) and CS2 (17 μL, 0.28 mmol) were added to a

solution of AM9025 (80 mg, 0.23 mmol ) in anhydrous THF (5 mL). The reaction was allowed

to stir under argon for 48 hours. The reaction mixture was then concentrated and

chromatographed on silica gel to yield 21, AM9029, (41 mg, 49%) as an oil. Rf = 0.40 (35%

ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.92 (m, 1 H) 5.52 -

75

5.66 (m, 1 H) 5.32 - 5.46 (m, 7 H) 4.05 (dd, J=5.1, 2.7 Hz, 2 H) 3.54 (t, J=6.6 Hz, 2 H) 2.78 -

2.88 (m, 6 H) 2.45 - 2.51 (m, 2 H) 2.19 - 2.25 (m, 3 H) 2.12 (q, J=6.8 Hz, 2 H) 1.73 (quin, J=7.6

Hz, 2 H). 13C NMR (126 MHz, CHLOROFORM-d) δ ppm 172.7, 132.3, 129.3, 129.0, 128.8,

128.6, 128.1, 127.7, 124.5, 79.9, 71.7, 45.0, 35.9, 29.3, 28.3, 26.7, 26.0, 25.9 (2C), 25.56. IR

(neat) cm-1 3294, 2186, 2092, 1647, 1535.

O

NH

NCS

(5Z,8Z,11Z,14Z)-18-isothiocyanato-N-(prop-2-ynyl)octadeca-5,8,11,14-tetraenamide, 22,

AM9039. Synthesized following the procedure for 21 from 18 (65 mg, 0.18 mmol) to yield 22,

AM9039 (65 mg, 96%) as a yellow oil. Rf = 0.34 (35% ethyl acetate / hexanes). 1H NMR (500

MHz, CHLOROFORM-d) δ ppm 5.71 (br. s., 1 H) 5.31 - 5.51 (m, 8 H) 4.06 (dd, J=4.9, 2.4 Hz,

2 H) 3.52 (t, J=6.8 Hz, 2 H) 2.71 - 2.89 (m, 6 H) 2.24 (t, J=2.9 Hz, 1 H) 2.17 - 2.22 (m, 2 H) 2.00

- 2.15 (m, 4 H) 1.66 - 1.80 (m, 4 H). 13C NMR (126 MHz, CHLOROFORM-d) δ 172.6, 130.3,

129.2, 129.2, 128.7, 128.5, 128.3, 128.3, 128.2, 79.9, 71.8, 45.3, 36.0, 31.8, 30.2, 29.4, 28.7,

26.8, 26.7, 25.9, 25.5. IR (neat) cm-1 3293, 2186, 2094, 1647, 1534.

O

NH

NCS

(5Z,8Z,11Z,14Z)-19-isothiocyanato-N-(prop-2-ynyl)nonadeca-5,8,11,14-tetraenamide, 23,


AM9017, (66 mg, 60%) as a yellow oil. Rf = 0.33 (35% ethyl acetate / hexanes). 1H NMR (500

MHz, CHLOROFORM-d) δ ppm 5.65 (br. s., 1 H) 5.32 - 5.48 (m, 8 H) 4.05 (dd, J=4.9, 2.4 Hz,

2 H) 3.53 (t, J=6.4 Hz, 2 H) 2.70 - 2.88 (m, 6 H) 2.23 (br. s., 1 H) 2.21 (t, J=7.8 Hz, 3 H) 2.06 -

76

2.17 (m, 4 H) 1.67 - 1.78 (m, 4 H) 1.50 (quin, J=7.6 Hz, 2 H). 13C NMR (126 MHz,


79.8, 71.7, 45.2, 35.9, 29.7, 29.3, 27.4, 26.8, 26.7, 26.5, 25.9 (2 C), 25.5. IR (neat) cm-1 3297,

2183, 2091, 1647, 1535.

O

NH

SCN

(5Z,8Z,11Z,14Z)-20-isothiocyanato-N-(prop-2-ynyl)icosa-5,8,11,14-tetraenamide, 24,


AM9004, (19mg, 80%) as a yellow oil. Rf = .30 (35% ethyl acetate/hexane). 1H NMR (500

MHz, CHLOROFORM-d) δ ppm 5.32 - 5.44 (m, 8 H) 4.05 (dd, J=5.4, 2.44 Hz, 2 H) 3.52 (t,

J=6.6 Hz, 2 H) 2.75 - 2.88 (m, 6 H) 2.23 (t, J=2.44 Hz, 1 H) 2.19 (t, J=7.5 Hz, 2 H) 2.05 - 2.16

(m, 4 H) 1.73 (quin, J=7.3 Hz, 2 H) 1.60 (quin, J=6.5 Hz, 2 H) 1.36 - 1.44. IR (neat) cm-1 3295,

2183, 2092, 1646, 1535. 13C NMR (126 MHz, CHLOROFORM-d) δ ppm 172.5, 130.3, 129.4,

129.3, 129.1, 129.0, 128.7, 128.5, 128.4, 79.7, 71.8, 45.0, 35.8, 29.6, 29.4, 27.7, 26.8, 26.7, 26.4,

25.9 (2 C), 25.7, 25.5. HRMS for C24H35N2OS (MH+) 399.2460. Calcd. 399.2470.

OH

hex-5-yn-1-ol, 28. A solution of 5-hexynoic acid (3.0 mL, 27.2 mmol) in anhydrous Et2O (15

mL) was slowly added to a 1.0M solution of LiAlH4 in Ether (27.2 mL, 27.2 mmol) at 0oC. The

reaction mixture was stirred under argon as it was warmed to room temperature and stirred for an

additional hour. A 1.0M HCl solution was added drop wise until the reaction mixture was

acidic. The lipophilic products were extracted with Et2O, and the aqueous layer was washed

77

with Et2O three additional times. The combined layers were dried with MgSO4 and concentrated

to give 5-hexyn-1-ol (2.09g, 78%) as an oil. Rf = 0.31 (35% ethyl acetate / hexanes). 1H NMR

(500 MHz, CHLOROFORM-d) δ ppm 3.69 (t, J=5.9 Hz, 2 H) 2.25 (td, J=6.8, 2.4 Hz, 2 H) 1.96

(t, J=2.4 Hz, 1 H) 1.70 (quin, J=6.4 Hz, 2 H) 1.63 (quin, J=7.3 Hz, 2 H).

OH

hept-6-yn-1-ol, 29. Synthesized following the procedure for 28, (3.49 g, 97%) colorless oil. Rf =

0.30 (30% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ 3.65 - 3.71 (m,

2H), 2.23 (dt, J = 2.7, 7.0 Hz, 2H), 1.97 (t, J = 2.7 Hz, 2H), 1.59 - 1.65 (m, 4H), 1.51 - 1.54 (m,

2H).

TMS OH

7-(trimethylsilyl)hepta-3,6-diyn-1-ol, 30. (3-bromoprop-1-ynyl)trimethylsilane 25 (1.15mL, 7.2

mmol) and 3-butyn-1-ol 26 (545 μL, 7.2 mmol) were added to a suspension of CuI (2.74 g, 14.4

mmol), NaI (2.15 g, 14.4 mmol), and K2CO3 (1.99 g, 14.4 mmol) in anhydrous DMF (7mL )

under an atmosphere of argon. The mixture was stirred at room temperature over night. The

mixture was then quenched with saturated a saturated NH4Cl solution. The lipophilic product

was extracted with Et2O, filtered, and washed with water, brine, and dried over MgSO4. The

organic layer was concentrated and the residue was chromatographed on silica gel to yield 30

(1.05 g, 70%) as yellow oil. Rf = 0.47 (35% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 3.64 - 3.76 (m, 2 H) 3.21 (t, J=2.4 Hz, 2 H) 2.43 - 2.49 (m, 2 H).

TMSOH

8-(trimethylsilyl)octa-4,7-diyn-1-ol, 31. Synthesized following the procedure for 30 from 25

(1.15 mL, 7.2 mmol) and 27 (670 μL, 7.2 mmol) to yield 31 (881 mg, 63%) as a yellow oil. Rf =

78

0.52 (35% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.76 (t,

J=6.1 Hz, 2 H) 3.19 (t, J=2.2 Hz, 2 H) 2.30 (tt, J=6.8, 4.7, 1.9 Hz, 2 H) 1.76 (quin, J=6.5 Hz, 2

H) 0.12 - 0.20 (m, 9 H).

TMS OH

9-(trimethylsilyl)nona-5,8-diyn-1-ol, 32. Synthesized following the procedure for 30 from 25

(1.15 mL, 7.2 mmol) and 28 (780 μL, 7.2 mmol) to yield 32 (816 mg, 54%) as a yellow oil. Rf =

0.51 (40% acetone / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.64 - 3.71 (m, 2

H) 3.19 (t, J=2.4 Hz, 2 H) 2.22 (tt, J=6.8, 2.3 Hz, 2 H) 1.64 - 1.71 (m, 2 H) 1.56 - 1.62 (m, 2 H).

TMSOH

10-(trimethylsilyl)deca-6,9-diyn-1-ol, 33. Synthesized following the procedure for 30 from 25

(1.00 mL, 6.2 mmol) and 29 (780 μL, 6.2 mmol) to yield 33 (1.00 g, 73%) as a yellow oil. Rf =

0.37 (35% ethyl acetate/hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.65 (br. s., 2

H) 3.19 (t, J=2.4 Hz, 2 H) 2.19 (tt, J=6.8, 2.4 Hz, 2 H) 1.57 - 1.62 (m, 2 H) 1.50 - 1.56 (m, 2 H)

1.43 - 1.49 (m, 2 H).

HOO

O

methyl 10-hydroxydeca-5,8-diynoate, 36. Synthesized following the procedure for 30 from 4-

chlorobut-2-yn-1-ol 34 (580 μL, 6.7 mmol) and methyl hex-5-ynoate 35 (.88mL, 13.4mmol) to

yield 36 (965 mg, 74%) as a colorless oil. Rf = 0.36 (35% ethyl acetate/hexane). 1H NMR (500

MHz, CHLOROFORM-d) δ ppm 4.27 (m, 2 H) 3.69 (s, 3 H) 3.18 (quin, J=2.2 Hz, 2 H) 2.44 (t,

J=7.6 Hz, 2 H) 2.24 (tt, J=6.8, 2.44 Hz, 2 H) 1.83 (quin, J=7.2 Hz, 2 H) 1.56 (br. s., 1 H).

BrO

O

79

methyl 10-bromodeca-5,8-diynoate, 37. A solution of PPh3 (1.44 g, 5.47 mmol) in anhydrous

CH2Cl2 (10 mL) was added drop wise to a stirred suspension of CBr4 (1.81 g, 5.47 mmol) and 36

(965 mg, 4.97 mmol) in CH2Cl2 (20mL) at 0oC. After 2 h the solvent was removed under reduced

pressure and the resulting residue was chromatographed on silica gel to yield 37 (1.09 g, 85%) as

a yellow oil. Rf = 0.62 (35% ethyl acetate/hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ

ppm 3.92 (t, J=2.2 Hz, 2 H) 3.68 (s, 3 H) 3.21 (t, J=2.2 Hz, 2 H) 2.44 (t, J=7.6 Hz, 2 H) 2.24 (tt,

J=6.8, 2.4 Hz, 2 H) 1.82 (quin, J=7.2 Hz, 3 H).

O

O

HO

methyl 17-hydroxyheptadeca-5,8,11,14-tetraynoate, 38. A solution of 37 (1.38 g, 5.39 mmol)

and 30 (971 mg, 5.39 mmol) in anhydrous DMF (6 mL) and 1.0M TBAF (5.39 g, 5.39 mmol) in

THF were added to a suspension of CuI (2.06 g, 10.8 mmol), NaI (1.62 g, 10.8 mmol), K2CO3

(1.49 g, 10.8 mmol) anhydrous DMF (5 mL). The reaction mixture was allowed to stir at room

temperature under argon for 72h. The reaction mixture was quenched with a saturated NH4Cl

solution and the lipophilic products were extracted with Et2O; washed with water, brine, and

dried over MgSO4. The ethereal layer was removed under reduced pressure and

chromatographed on silica gel to yield methyl 38 (690 mg, 45%) as a yellow oil. Rf = 0.19 (35%

ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.71 (q, J=6.0 Hz, 2 H)

3.68 (s, 3 H) 3.12 - 3.18 (m, 6 H) 2.41 - 2.48 (m, 4 H) 2.24 (tt, J=7.0, 2.3 Hz, 2 H) 1.82 (quin,

J=7.2 Hz, 2 H).

O

OHO

methyl 18-hydroxyoctadeca-5,8,11,14-tetraynoate, 39. Synthesized following the procedure

for 38 from 37 (1.44 g, 5.6 mmol) and 31 (881 mg, 4.5 mmol) to yield 39 (455 mg, 34%) as a

80

yellow oil. Rf = 0.13 (35% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ

ppm 3.75 (q, J=5.8 Hz, 2 H) 3.68 (s, 3 H) 3.09 - 3.20 (m, 6 H) 2.43 (t, J=7.5 Hz, 3 H) 2.30 (m, 2

H) 2.21 - 2.27 (m, 2 H) 1.82 (quin, J=7.2 Hz, 2 H) 1.75 (quin, J=6.6 Hz, 2 H).

O

O

HO

methyl 19-hydroxynonadeca-5,8,11,14-tetraynoate, 40. Synthesized following the procedure

for 38 from 37 (962 mg, 3.74 mmol) and 32 (780 mg, 3.74 mmol) to yield 40 (512 mg, 63%) as a

yellow oil. Rf = 0.17 (35% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ

ppm 3.68 (s, 3 H) 3.65 - 3.67 (m, 2 H) 3.09 - 3.19 (m, 6 H) 2.43 (t, J=7.3 Hz, 2 H) 2.18 - 2.28

(m, 4 H) 1.82 (quin, J=7.2 Hz, 2 H) 1.67 (quin, J=6.8 Hz, 2 H) 1.59 (quin, J=7.3 Hz, 2 H).

O

OHO

methyl 20-hydroxyicosa-5,8,11,14-tetraynoate, 41. Synthesized following the procedure for 38

from 37 (900 mg, 3.5 mmol) and 33 (778 mg, 3.5 mmol) to yield 41 (605 mg, 53%) as a yellow

oil. Rf = 0.16 (35% ethyl acetate/hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 3.68

(s, 3 H) 3.65 (t, 2 H) 3.10 - 3.19 (m, 6 H) 2.43 (t, J=7.3 Hz, 2 H) 2.24 (tt, J=6.7, 4.5 Hz, 2 H)

2.18 (tt, J=4.6, 2.3 Hz, 2 H) 1.82 (quin, J=7.2 Hz, 2 H) 1.58 (quin, 2 H) 1.49 - 1.55 (m, 2 H) 1.42

- 1.49 (m, 2 H).

O

OOH

(5Z,8Z,11Z,14Z)-methyl 17-hydroxyheptadeca-5,8,11,14-tetraenoate, 42. To a solution of

Ni(OAc)2•4H2O (1.03 g, 4.13 mmol) in anhydrous methanol (30 mL), was added NaBH4 (183

mg, 4.86 mmol) at room temperature under and atmosphere of argon. This mixture was

immediately put under vacuum and purged with H2 (3 times) and allowed to stir for 5 minutes.

81

This solution was treated with ethylenediamine (422 μL, 6.32 mmol) and stirred for an additional

5 minutes; at which 38 (690 mg, 2.43 mmol) in anhydrous methanol (20 mL) was added. The

mixture was stirred at room temp under H2 for 2h. The reaction mixture was filtered through

celite, where the filtrate was diluted with Et2O and brine. The organic phase was separated and

the aqueous phase was extracted 3 times with Et2O, and the combined organic layers were dried

over MgSO4. The ethereal layer was removed under reduced pressure and the resulting residue

was chromatographed on silica gel to yield 42 (237 mg, 33%) as a colorless oil. Rf = 0.28 (35%

ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.48 - 5.62 (m, 1 H)

5.27 - 5.48 (m, 7 H) 3.54 - 3.77 (m, 5 H) 2.63 - 2.98 (m, 6 H) 2.38 (q, J=6.5 Hz, 2 H) 2.33 (t,

J=7.3 Hz, 2 H) 2.12 (q, J=6.8 Hz, 2 H) 1.71 (quin, J=7.5 Hz, 2 H).

O

O

OH

(5Z,8Z,11Z,14Z)-methyl 18-hydroxyoctadeca-5,8,11,14-tetraenoate, 43. Synthesized

following the procedure for 42 from 39 (455 mg, 1.52 mmol) to yield 43 (343 mg, 56%) as a

colorless oil. Rf = 0.13 (35% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ

ppm 5.31 - 5.46 (m, 8 H) 3.63 - 3.69 (m, 5 H) 2.75 - 2.87 (m, 6 H) 2.33 (t, J=7.6 Hz, 2 H) 2.17

(q, J=7.0 Hz, 2 H) 2.11 (q, J=7.0 Hz, 2 H) 1.70 (quin, J=7.6 Hz, 2 H) 1.64 (quin, J=7.3 Hz, 2 H).

O

O

OH

(5Z,8Z,11Z,14Z)-methyl 19-hydroxynonadeca-5,8,11,14-tetraenoate, 44. Synthesized

following the procedure for 42 from 40 (512 mg, 1.64 mmol) to yield 44 (250 mg, 48%) as a

colorless oil. Rf = 0.52 (35% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ

82

ppm 5.31 - 5.45 (m, 8 H) 3.67 (s, 3 H) 3.64 (t, J=6.1 Hz, 2 H) 2.74 - 2.87 (m, 6 H) 2.33 (t, J=7.6

Hz, 2 H) 2.05 - 2.15 (m, 4 H) 1.71 (quin, J=7.5 Hz, 2 H) 1.56 - 1.64 (m, 2 H) 1.45 (quin, J=6.8

Hz, 2 H).

O

O

HO

(5Z,8Z,11Z,14Z)-methyl 20-hydroxyicosa-5,8,11,14-tetraenoate, 45. Synthesized following

the procedure for 42 from 41 (605 mg, 1.85 mmol) to yield 45 (354 mg, 57%) as a colorless oil.

Rf = 0.49 (35% ethyl acetate/hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.30 -

5.47 (m, 8 H) 3.67 (s, 3 H) 3.64 (t, J=6.1 Hz, 2 H) 2.74 - 2.87 (m, 6 H) 2.33 (t, J=7.3 Hz, 2 H)

2.05 - 2.15 (m, 4 H) 1.71 (quin, J=7.6 Hz, 2 H) 1.57 (quin, J=6.4 Hz, 2 H) 1.34 - 1.44 (m, 4 H).

O

ON3

(5Z,8Z,11Z,14Z)-methyl 17-azidoheptadeca-5,8,11,14-tetraenoate, 46. DBU (182 μL, 1.22

mmol) and DPPA (263 μL, 1.22 mmol) were added to a solution of 42 (237 mg, 0.81 mmol) in

anhydrous DMF (3 mL) at 120oC. The reaction was allowed to stir for 4h where up it was

diluted with ether, washed with water, brine, and dried of MgSO4. The ethereal layer was

evaporated off under reduced pressure and the resulting residue was chromatographed on silica

to yield 46 (134 mg, 52%) as a yellow oil. Rf = 0.83 (35% ethyl acetate / hexanes). 1H NMR

(500 MHz, CHLOROFORM-d) δ ppm 5.48 - 5.56 (m, 1 H) 5.32 - 5.45 (m, 7 H) 3.67 (s, 3 H)

3.30 (t, J=7.1 Hz, 2 H) 2.74 - 2.88 (m, 6 H) 2.39 (q, J=7.3 Hz, 2 H) 2.33 (t, J=7.3 Hz, 2 H) 2.08 -

2.15 (m, 2 H) 1.71 (quin, J=7.5 Hz, 2 H).

83

O

O

N3

(5Z,8Z,11Z,14Z)-methyl 18-azidooctadeca-5,8,11,14-tetraenoate, 47. Synthesized following

the procedure for 46 from 43 (340 mg, 1.11 mmol) to yield 47 (93 mg, 25%) as a yellow oil. Rf =

0.75 (35% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.31 - 5.48

(m, 8 H) 3.67 (s, 3 H) 3.27 (m, J=6.8 Hz, 2 H) 2.72 - 2.87 (m, 6 H) 2.32 (t, J=7.3 Hz, 3 H) 2.17

(q, J=6.8 Hz, 1 H) 2.11 (q, J=6.8 Hz, 2 H) 1.63 - 1.75 (m, 4 H).

O

O

N3

(5Z,8Z,11Z,14Z)-methyl 19-azidononadeca-5,8,11,14-tetraenoate, 48. Synthesized following

the procedure for 46 from 44 (472 mg, 1.47 mmol) to yield 48 (228 mg, 45%) as a yellow oil. Rf

= 0.83 (35% ethyl acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.30 -

5.45 (m, 8 H) 3.67 (s, 3 H) 3.27 (t, J=6.8 Hz, 2 H) 2.81 (d, 5 H) 2.32 (t, J=7.3 Hz, 2 H) 2.02 -

2.15 (m, 4 H) 1.71 (quin, J=7.5 Hz, 2 H) 1.62 (quin, J=7.3 Hz, 2 H) 1.45 (quin, J=7.5 Hz, 2 H).

O

O

N3

(5Z,8Z,11Z,14Z)-methyl 20-azidoicosa-5,8,11,14-tetraenoate, 49. Synthesized following the

procedure for 46 from 45 (77 mg, 0.23 mmol) to yield 49 (45 mg, 54%) as a yellow oil. Rf = 0.86

(35% ethyl acetate/hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.32 - 5.44 (m, 8

H) 3.67 (s, 3 H) 3.26 (t, J=6.8 Hz, 2 H) 2.74 - 2.87 (m, 6 H) 2.32 (t, J=7.5 Hz, 2 H) 2.02 - 2.15

(m, 4 H) 1.71 (quin, J=7.5 Hz, 2 H) 1.61 (quin, J=6.4 Hz, 2 H) 1.35 - 1.43 (m, 4 H).

84

O

OHN3

(5Z,8Z,11Z,14Z)-17-azidoheptadeca-5,8,11,14-tetraenoic acid, 50. A 1.0M solution of LiOH

(84 μL, 0.84 mmol) was added to a solution of 46 (134 mg, 0.42 mmol) in THF (5 mL). The

reaction was allowed to stir under argon for 48 hours. Upon completion, a 1.0M solution of HCl

was added until the reaction mixture was slightly acidic. The lipophilic products were extracted

with Et2O washed with brine, and dried over MgSO4. The ethereal layer was evaporated off

under reduced pressure to yield 50 (102 mg, 80%) as yellow oil. Rf = 0.28 (35% ethyl acetate /

hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.48 - 5.58 (m, 1 H) 5.32 - 5.46 (m, 7

H) 3.30 (t, J=6.8 Hz, 2 H) 2.74 - 2.88 (m, 6 H) 2.33 - 2.43 (m, 4 H) 2.14 (q, J=7.2 Hz, 2 H) 1.72

(quin, J=7.5 Hz, 2 H).

O

OH

N3

(5Z,8Z,11Z,14Z)-18-azidooctadeca-5,8,11,14-tetraenoic acid, 51. Synthesized following the

procedure for 50 from 47 (100 mg, 0.31 mmol) to yield 51 (46 mg, 45%) as a yellow oil. Rf =


(m, 8 H) 3.19 - 3.33 (m, 2 H) 2.71 - 2.91 (m, 6 H) 2.37 (t, J=7.3 Hz, 2 H) 2.08 - 2.20 (m, 4 H)

1.63 - 1.76 (m, 4 H).

O

OH

N3

(5Z,8Z,11Z,14Z)-19-azidononadeca-5,8,11,14-tetraenoic acid, 52. Synthesized following the

procedure for 50 from 48 (228 mg, 0.66 mmol) to yield 52 (166 mg, 76%) as a yellow oil. Rf =

85


(m, 8 H) 3.27 (t, J=6.8 Hz, 2 H) 2.74 - 2.88 (m, 6 H) 2.37 (t, J=7.3 Hz, 2 H) 2.06 - 2.17 (m, 4 H)

1.72 (quin, J=7.5 Hz, 2 H) 1.62 (quin, J=6.8 Hz, 2 H) 1.45 (quin, J=7.8 Hz, 2 H).

O

OH

N3

(5Z,8Z,11Z,14Z)-20-azidoicosa-5,8,11,14-tetraenoic acid, 53. Synthesized following the

procedure for 50 from 49 (45 mg, 0.12 mmol) to yield 53 (32 mg, 76%) as a yellow oil. Rf = 0.22

(35% ethyl acetate/hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.33 - 5.46 (m, 8 H)

3.27 (t, J=6.8 Hz, 2 H) 2.76 - 2.87 (m, 6 H) 2.37 (t, J=7.3 Hz, 2 H) 2.07 - 2.19 (m, 4 H) 1.73

(quin, J=7.5 Hz, 2 H) 1.61 (quin, J=6.4 Hz, 2 H) 1.36 - 1.43 (m, 4 H).

NH

O

O

(R)-tert-butyl but-3-yn-2-ylcarbamate, 58. Oxalylchloride (498 μL, 5.8 mmol) was dissolved

in anhydrous CH2Cl2 (25 mL), stirred and cooled to -78 °C, under an argon atmosphere. This

solution was treated with DMSO (824 μL, 11.6 mmol) and stirred for 15 minutes. 54 (500 mg,

2.9 mmol) was added to the solution which stirred for 1.5h. The solution was then treated with

Et3N (2.42 mL, 17.4 mmol) and slowly warmed to room temperature. The solution was diluted

with ethyl acetate, washed with water, brine, and dried over MgSO4. The solvent was removed

under reduced pressure. The resulting aldehyde, 56, was dissolved in anhydrous MeOH (20 mL)

and treated with dimethyl 1-diazo-2-oxopropylphosphonate (836 mg, 4.4 mmol) and K2CO3

(801 mg, 5.8 mmol), stirred for 1 hour at 0° C and then overnight at room temperature. The

86

reaction was quenched with a saturated solution of NH4Cl. The methanol was removed under

reduced pressure and the product was extracted with ethyl acetate (3x) and dried with MgSO4.

The organic layer was removed under reduced pressure and the residue was chromatographed on

silica gel to yield 58 (273 mg, 56%) as a white solid. Rf = 0.53 (20% ethyl acetate / hexanes). 1H

NMR (500 MHz, CHLOROFORM-d) δ ppm 4.58 - 4.79 (m, 1 H) 4.39 - 4.56 (m, 1 H) 2.25 (d,

J=1.9 Hz, 1 H) 1.45 (s, 9 H) 1.40 (d, J=6.8 Hz, 3 H).

NH

O

O

(S)-tert-butyl but-3-yn-2-ylcarbamate, 59.

Synthesized following the procedure for 58 from 55 (500 mg, 2.9 mmol) to yield 59 (147 mg,

30%) as a white solid. Rf = 0.53 (20% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 4.61 - 4.75 (m, 1 H) 4.41 - 4.56 (m, 1 H) 2.25 (d, J=2.4 Hz, 1 H) 1.45

(s, 9 H) 1.40 (d, J=6.8 Hz, 3 H).

NH3 O

O

CF3

(R)-but-3-yn-2-aminium 2,2,2-trifluoroacetate, 60. A solution of 58 (140 mg, 0.83 mmol) in

CH2Cl2 (4 mL) was treated with trifluoroacetic acid (1 mL) at room temperature and stirred for

2h. Solvent and excess trifluoroacetic acid was removed under reduced pressure, resulting in 60

(120 mg, 86%) as an oil. 1H NMR (500 MHz, WATER-d2) δ ppm 4.74 (dq, J=2.4, 6.8 Hz, 1 H)

3.47 (d, J=2.4 Hz, 1 H) 2.05 (d, J=6.8 Hz, 3 H).

NH3 O

O

CF3

87

(S)-but-3-yn-2-aminium 2,2,2-trifluoroacetate, 61. Synthesized following the procedure for 60

from 59 (140 mg, 0.83 mmol) to yield 61 (140 mg, 90%) as an oil. 1H NMR (500 MHz, Water-

d2) δ ppm 4.77 (q, J=6.8 Hz, 1 H) 3.50 (t, J=2.4 Hz, 1 H) 2.08 (d, J=7.3 Hz, 3 H).

O

NH

N3

(5Z,8Z,11Z,14Z)-20-azido-N-((R)-but-3-yn-2-yl)icosa-5,8,11,14-tetraenamide, 62, AM9069.

EDCI (115 mg, 0.60 mmol), DMAP (3 mg, 0.03 mmol), triethylamine (0.024 mL, 0.17 mmol)

and 60 (30 mg, 0.17 mmol) were added to a solution of 52 (47 mg, 0.14 mmol) in anhydrous

DCM (5 mL) at 0°C. The reaction was allowed to stir under argon for 4 hours. Upon completion

the reaction was diluted with CH2Cl2 and water. The organic layer was separated and

concentrated. The resulting residue was chromatographed on silica gel to yield 62, AM9069, (56

mg, 100%) as a yellow oil. Rf = 0.56 (35% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 5.74 (br. s., 1 H) 5.31 - 5.45 (m, 8 H) 4.79 - 4.87 (m, 1 H) 3.26 (t,

J=7.1 Hz, 2 H) 2.74 - 2.87 (m, 6 H) 2.25 (d, J=2.4 Hz, 1 H) 2.17 (t, J=7.3 Hz, 2 H) 1.72 (quin,

J=7.3 Hz, 2 H) 1.57 - 1.64 (m, 2 H) 1.40 (d, J=6.8 Hz, 3 H) 1.36 (dd, J=6.8, 4.4 Hz, 2 H) 1.27 -

1.34 (m, 4 H). 13C NMR (126 MHz, CHLOROFORM-d) δ ppm 171.8, 130.1, 129.8, 129.3,

129.0, 128.6, 128.4, 128.3, 128.2, 84.5, 70.5, 51.6, 37.0, 36.0, 31.8, 29.3, 29.0, 27.4, 27.3, 26.8,

26.6, 25.9, 22.5, 21.2. IR (neat) cm-1 3302, 2094, 1643, 1535, 1453.

O

NH

N3

88

(5Z,8Z,11Z,14Z)-20-azido-N-((S)-but-3-yn-2-yl)icosa-5,8,11,14-tetraenamide, 63, AM9073.



CHLOROFORM-d) δ ppm 5.65 (br. s., 1 H) 5.30 - 5.45 (m, 8 H) 4.78 - 4.87 (m, 1 H) 3.26 (t,

J=6.8 Hz, 2 H) 2.63 - 2.89 (m, 6 H) 2.25 (d, J=2.4 Hz, 1 H) 2.17 (t, J=7.8 Hz, 2 H) 1.94 - 2.13

(m, 4 H) 1.71 (quin, J=7.3 Hz, 2 H) 1.61 (quin, J=6.8 Hz, 2 H) 1.41 (d, J=6.8 Hz, 3 H) 1.34 -

1.39 (m, 4 H). 13C NMR (100 MHz, CHLOROFORM-d) δ ppm 171.9, 130.1, 129.2, 129.0,

128.6, 128.4, 128.3, 128.25, 128.20, 84.4, 70.5, 51.6, 36.9, 36.1, 31.8, 31.6, 29.3, 28.8, 27.3,

26.8, 26.5, 25.7, 22.5, 21.3. IR (neat) cm-1 3305, 2094, 1644, 1536, 1454.

O

NH

SCN

(5Z,8Z,11Z,14Z)-N-((R)-but-3-yn-2-yl)-20-isothiocyanatoicosa-5,8,11,14-tetraenamide, 64,


AM9070, (19 mg, 45%) as a yellow oil. Rf = 0.67 (35% ethyl acetate / hexanes). 1H NMR (500

MHz, CHLOROFORM-d) δ ppm 5.62 (br. s., 1 H) 5.22 - 5.47 (m, 8 H) 4.82 (m, 1 H) 3.51 (t,

J=6.6 Hz, 2 H) 2.73 - 2.88 (m, 6 H) 2.25 (d, J=1.9 Hz, 1 H) 2.17 (t, J=7.3 Hz, 2 H) 2.06 - 2.15

(m, 4 H) 1.67 - 1.77 (m, 4 H) 1.41 (d, J=6.8 Hz, 3 H) 1.28 - 1.34 (m, 4 H). 13C NMR (100 MHz,

CHLOROFORM-d) δ 171.6, 131.1, 123.0, 129.5, 128.9, 128.7, 128.4, 128.3, 128.2, 84.4, 70.8,

45.3, 36.9, 36.1, 30.5, 29.7, 29.2, 27.6, 27.4, 26.7, 26.3, 26.0, 25.6, 22.5. IR (neat) cm-1 3293,

2185, 2094, 1645, 1537, 1452.

89

O

NH

SCN

(5Z,8Z,11Z,14Z)-N-((S)-but-3-yn-2-yl)-20-isothiocyanatoicosa-5,8,11,14-tetraenamide, 65,


AM9074, (16 mg, 52%) as an oil. Rf = 0.59 (35% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 5.63 (br. s., 1 H) 5.30 - 5.44 (m, 8 H) 4.83 (m, 1 H) 3.52 (t, J=6.6 Hz,

2 H) 2.71 - 2.88 (m, 6 H) 2.26 (d, J=2.4 Hz, 1 H) 2.17 (t, J=8.3 Hz, 2 H) 2.05 - 2.14 (m, 4 H)

1.67 - 1.76 (m, 4 H) 1.42 - 1.47 (m, 2 H) 1.41 (d, J=6.8 Hz, 3 H) 1.34 - 1.39 (m, 2 H). 13C NMR

13C NMR (100 MHz, CHLOROFORM-d) δ 171.8, 131.1, 129.9, 129.3, 129.0, 128.6, 128.4,

128.4, 128.3, 84.5, 70.6, 45.2, 37.0, 36.1, 30.1, 29.5, 29.0, 27.4, 27.2, 26.8, 26.4, 25.9, 25.5, 22.6.

IR (neat) cm-1 3295, 2184, 2094, 1644, 1536, 1452.

90

2.13 References

1. Baker, B. R. Design of active-site directed irreversible enzyme inhibitors; the organic chemistry of the enzymic active-site [by] B. R. Baker. New York, Wiley [1967]: 1967. 2. Vallee, B. L.; Riordan, J. F. Chemical approaches to the properties of active sites of enzymes. Annu. Rev. Biochem. 1969, 38, 733-4. 3. Knowles, J. R. Photogenerated reagents for biological receptor-site labeling. Accounts of Chemical Research 1972, 5, 155-160. 4. Pei, Y.; Mercier, R. W.; Anday, J. K.; Thakur, G. A.; Zvonok, A. M.; Hurst, D.; Reggio, P. H.; Janero, D. R.; Makriyannis, A. Ligand-Binding Architecture of Human CB2 Cannabinoid Receptor: Evidence for Receptor Subtype-Specific Binding Motif and Modeling GPCR Activation. Chemistry & Biology 2008, 15, 1207-1219. 5. Picone, R. P.; Fournier, D. J.; Makriyannis, A. Ligand based structural studies of the CB1 cannabinoid receptor. Journal of Peptide Research 2002, 60, 348-356. 6. Zvonok, N.; Xu, W.; Williams, J.; Janero, D. R.; Krishnan, S. C.; Makriyannis, A. Mass Spectrometry-Based GPCR Proteomics: Comprehensive Characterization of the Human Cannabinoid 1 Receptor. Journal of Proteome Research 2010, 9, 1746-1753. 7. Charalambous, A.; Yan, G.; Houston, D. B.; Howlett, A. C.; Compton, D. R.; Martin, B. R.; Makriyannis, A. 5'-Azido-A8-THC: A Novel Photoaffinity Label for the Cannabinoid Receptor. Journal of Medicinal Chemistry 1992, 35, 3076-3079. 8. Guo, Y.; Abadji, V.; Morse, K. L.; Fournier, D. J.; Li, X.; Makriyannis, A. (-)-1 l-Hydroxy-7’-isothiocyanato-l1’’-,dimethylheptyl-A8-THC:A Novel,High-Affinity Irreversible Probe for the Cannabinoid Receptor in the Brain. Journal of Medicinal Chemistry 1994, 37, 3867-3870. 9. Morse, K. L.; Fournier, D. J.; Li, X.; Grzybowska, J.; Makriyannis, A. A novel electrophilic high affinity irreversible probe for the cannabinoid receptor. Life Sciences 1995, 56, 1957-1962. 10. Picone, R. P.; Khanolkar, A. D.; Xu, W.; Ayotte, L. A.; Thakur, G. A.; Hurst, D. P.; Abood, M. E.; Reggio, P. H.; Fournier, D. J.; Makriyannis, A. (-)-7′-Isothiocyanato-11-hydroxy-1′,1′-dimethylheptylhexahydrocannabinol (AM841), a High-Affinity Electrophilic Ligand, Interacts Covalently with a Cysteine in Helix Six and Activates the CB1 Cannabinoid Receptor. Molecular Pharmacology 2005, 68, 1623-1635. 11. Balas, L.; Cascio, M. G.; Marzo, V. D.; Durand, T. Synthesis of a potential photoactivatable anandamide analog. Bioorganic & Medicinal Chemistry Letters 2006, 16, 3765-3768. 12. Martín-Couce, L.; Martín-Fontecha, M.; Capolicchio, S.; López-Rodríguez, M. a. L.; Ortega-Gutiérrez, S. Development of Endocannabinoid-Based Chemical Probes for the Study of Cannabinoid Receptors. Journal of Medicinal Chemistry 2011, 54, 5265-5269. 13. Li, C.; Xu, W.; Vadivel, S. K.; Fan, P.; Makriyannis, A. High Affinity Electrophilic and Photoactivatable Covalent Endocannabinoid Probes for the CB1 receptor. Journal of Medicinal Chemistry 2005, 48, 6423-6429. 14. Lin, S.; Khanolkar, A. D.; Fen, P.; Goutopoulos, A.; Qin, C.; Papahadjis, D.; Makriyannis, A. Novel Analogues of Arachidonylethanolamide (Anandamide): Affinities for the CB1 and CB2 Cannabinoid Receptors and Metabolic Stability. Journal of Medicinal Chemistry 1998, 41, 5353-5361.

91

15. Tedeschi, C.; Saccavini, C.; Maurette, L.; Soleilhavoup, M. l.; Chauvin, R. 1,4-Diynes from alkynyl-propargyl coupling reactions. Journal of Organometallic Chemistry 2003, 670, 151-169. 16. Montel, F.; Beaudegnies, R.; Kessabi, J.; Martin, B.; Muller, E.; Wendeborn, S.; Jung, P. M. J. New Method Based on 1-(Trimethylsilyl)alk-1-yne To Prepare 1,4-Skipped Diynes. Organic Letters 2006, 8, 1905-1908. 17. Appel, R. Tertiary Phosphane/Tetrachloromethane, a Versatile Reagent for Chlorination, Dehydration, and P N Linkage. Angewandte Chemie International Edition in English 1975, 14, 801-811. 18. Lindlar, H. Ein neuer Katalysator für selektive Hydrierungen. Helvetica Chimica Acta 1952, 35, 446-450. 19. Brown, C. A. Catalytic hydrogenation. V. Reaction of sodium borohydride with aqueous nickel salts. P-1 nickel boride, a convenient, highly active nickel hydrogenation catalyst. The Journal of Organic Chemistry 1970, 35, 1900-1904. 20. Yao, F.; Li, C.; Vadivel, S. K.; Bowman, A. L.; Makriyannis, A. Development of novel tail-modified anandamide analogs. Bioorganic & Medicinal Chemistry Letters 2008, 18, 5912-5915. 21. Abadji, V.; Lin, S.; Taha, G.; Griffin, G.; Stevenson, L. A.; Pertwee, R. G.; Makriyannis, A. (R)-Methanandamide: A Chiral Novel Anandamide Possessing Higher Potency and Metabolic Stability. Journal of Medicinal Chemistry 1994, 37, 1889-1893. 22. Omura, K.; Swern, D. Oxidation of alcohols by “activated” dimethyl sulfoxide. a preparative, steric and mechanistic study. Tetrahedron 1978, 34, 1651-1660. 23. Müller, S.; Liepold, B.; Roth, G. J.; Bestmann, H. J. An Improved One-pot Procedure for the Synthesis of Alkynes from Aldehydes. Synlett 1996, 1996, 521-522. 24. Goutopoulos, A.; Fan, P.; Khanolkar, A. D.; Xie, X.-Q.; Lin, S.; Makriyannis, A. Stereochemical Selectivity of Methanandamides for the CB1 and CB2 Cannabinoid Receptors and Their Metabolic Stability. Bioorganic & Medicinal Chemistry 2001, 9, 1673-1684. 25. Urbani, P.; Cavallo, P.; Cascio, M. G.; Buonerba, M.; De Martino, G.; Di Marzo, V.; Saturnino, C. New metabolically stable fatty acid amide ligands of cannabinoid receptors: Synthesis and receptor affinity studies. Bioorganic & Medicinal Chemistry Letters 2006, 16, 138-141. 26. Hoegberg, T.; Stroem, P.; Ebner, M.; Raemsby, S. Cyanide as an efficient and mild catalyst in the aminolysis of esters. Journal of Organic Chemistry 1987, 52, 2033-2036. 27. Glass, M.; Faull, R. L. M.; Dragunow, M. Cannabinoid receptors in the human brain: a detailed anatomical and quantitative autoradiographic study in the fetal, neonatal and adult human brain. Neuroscience 1997, 77, 299-318. 28. Palczewski, K.; Kumasaka, T.; Hori, T.; Behnke, C. A.; Motoshima, H.; Fox, B. A.; Le Trong, I.; Teller, D. C.; Okada, T.; Stenkamp, R. E.; Yamamoto, M.; Miyano, M. Crystal Structure of Rhodopsin: A G Protein-Coupled Receptor. Science 2000, 289, 739. 29. Gnanaprakasam, B.; Milstein, D. Synthesis of amides from esters and amines with liberation of H2 under neutral conditions. J. Am. Chem. Soc. 2011, 133, 1682-1685. 30. Tsou, K.; Mackie, K.; Sañudo-Peña, M. C.; Walker, J. M. Cannabinoid CB1 receptors are localized primarily on cholecystokinin-containing GABAergic interneurons in the rat hippocampal formation. Neuroscience 1999, 93, 969-975. 31. Bezuglov, V.; Bobrov, M.; Gretskaya, N.; Gonchar, A.; Zinchenko, G.; Melck, D.; Bisogno, T.; Di Marzo, V.; Kuklev, D.; Rossi, J.-C.; Vidal, J.-P.; Durand, T. Synthesis and

92

biological evaluation of novel amides of polyunsaturated fatty acids with dopamine. Bioorganic & Medicinal Chemistry Letters 2001, 11, 447-449. 32. El Fangour, S.; Balas, L.; Rossi, J.-C.; Fedenyuk, A.; Gretskaya, N.; Bobrov, M.; Bezuglov, V.; Hillard, C. J.; Durand, T. Hemisynthesis and preliminary evaluation of novel endocannabinoid analogues. Bioorganic & Medicinal Chemistry Letters 2003, 13, 1977-1980.

93

CHAPTER 3

CHEMOENZYMATIC SYNTHESIS OF BIOLOGICALLY ACTIVE COMPOUNDS

94

3.1 Introduction

2-AG is an important endogenous signaling molecule that acts as an agonist at the

cannabinoid receptors. As a principal endocannabinoid, the investigation of 2-AG’s interaction

with the CB receptors and subsequent biological effects have been heavily studied,1-4 however,

chemical properties of this compound impede its study. 2-AG and other 2-monoacylglycerols

(2-MAGs) have a propensity to experience a thermodynamic acyl-migration from the sn-2 to the

more stable sn-1(3) position (Scheme 3.1).

Scheme 3.1 Acyl migration from 2-AG to 1(3)-AG

R O

OOH

OH R O

O

OHOH

12

3

12

3

Boswinkel, et al., reported kinetic experiments investigating the rate of acyl migration of

various fatty acids (butyryl, lauryl, caprylyl, and palmityl). They observed over a period of 168

hours, that the ratio of 1-MAG to 2-MAG ranged from 4.6 to 8.5. They also identified an

increase in the rate of acyl migration as the length of the acyl chain increased.5 Lyubachevskaya,

et al., observed an equilibrium of 1-MAG to 2-MAG of 7.3:1 over 30 hours at 37 °C in

chylomicra emulsions. Incubation of 2-MAGs at lower temperatures showed less acyl migration,

with no migration occurring at -10 °C.6

When 2-AG experiences acyl migration, the resulting 1-AG does not activate the

cannabinoid receptors in vitro.7 Obtaining large quantities of 2-AG can be difficult even through

synthesis. This becomes a major problem when trying to perform larger studies of 2-AG, as the

acyl migration is facile and encouraged by the presence of acid, base, heat, and protic solvents

95

commonly used for 2-AG synthesis.8, 9 To overcome these issues we have investigated moderate

reaction conditions utilizing enzymes to synthesize 2-AG, where minimal acyl migration is

observed.

3.1.1 Synthesis of 2-AG

There have been several reported syntheses of 2-AG, however, these employ many

laborious steps with unfavorable conditions and work-ups that may encourage acyl migration.

One of the earliest 2-AG syntheses reported by Han, et al., starts with the 1,3-triisopropylsilyl

(TIPS) protection of glycerol. The 1,3-TIPS glycerol is then coupled to arachidonic acid,

followed by removal of the silyl protecting groups over 24h with the use of TBAF and acetic

acid in 59% yield (Scheme 3.2).10

Scheme 3.2 TIPS method for synthesizing 2-AG

OHOH

HO

TIPSCl, DMF,imidazole

97%OTIPS

OHTIPSO

arachidonic acid,EDCI, DCM

94%

O

O

OTIPS

OTIPSO

O

OH

OH6eq AcOH, THF 6eq TBAF, -30 °C

59%

A second method coupled arachidonic acid with 1,3-benzylideneglycerol. Phenylboronic

acid is used to remove the benzylidene, resulting in a mixture of 1,3- and 2,3-phenylboronate

96

esters that required separation before subsequent boronate ester removal with methanol and

water in 86% yield (Scheme 3.3). 11

Scheme 3.3 2-AG synthesis through benzylidene

O

OH1) (COCl)2, DMF

2) HOO

O

H

Ph

O

OO

O Ph

H

PhB(OH)2

B(OEt)3

O

OO

BO

O

OPh

H2O, MeOH H2O, MeOH

O BO

Ph

2-AG 1-AG

The most recent method was developed by Stamatov, et al., starting from the glycidyl

ester of arachidonic acid that is opened with trifluoroacetic acid to produce a triacylglycerol. 2-

AG is then obtained after treatment with pyridine and methanol in 94% yield (Scheme 3.4).12

Scheme 3.4 2-AG synthesis through glycidal ring openeing

O

OO

O

O

O

O

O

CF3

O

CF3(F3CCO)2OPyridine, MeOH

2-AG

97

3.1.2 Lipase in the Synthesis of Acylated Glycerols

Application of lipase in the syntheses of 1,3-diacylglycerol, and 1(3)-rac-monoacyl

glycerol has been extensively studied and reviewed.13-21 The selectivity and yield are determined

by various factors which include the amount of enzyme, solvent, temperature and the type of

lipase used.22, 23 Even though the preparation of selective 1,3-diacylglycerols has been achieved

successfully, it has been a challenging task for the synthesis of 2-MAGs due to over hydrolysis

of the sn-2 acyl group and acyl migration from sn-2 to sn-1 or -3 position. Lipase-mediated

hydrolysis of triglycerides using 1,3-regiospecific lipases, esterification of fatty acids or

transesterification of fatty esters with glycerol, and the glycerolysis of triglycerides have been

documented.24, 25 Irimescu, et al., reported a successful synthesis of various 2-acylglycerols of

fatty acids through regiospecfic ethanolysis of symmetrical triglycerides with immobilized

Candida antarctica lipase (CAL, Novozyme 435).17, 26 Even though CAL is not considered as a

1,3-regiospecific enzyme, it has been consistently used for the preparation of 1,3-acylglycerols

and ethanolysis of triglycerides.26, 27 All existing methods utilize symmetrical (“AAA” type)

triglycerides which result in the formation of the corresponding ester by-product that requires

exhaustive purification (Scheme 3.5).

Scheme 3.5: “AAA” and “ABA” triglycerides

O

OCOR

OCORR

O

AAA typeO

OCOC3H7

OCOC3H7R

O

ABA type

R = fatty acid

Encouraged by this literature, we recently reported a method for the synthesis of 2-AG

that utilizes a structured 1,3-dibutyryl-2-arachidonate, an “ABA” type triglyceride. We chose

98

butyrate at the 1 and 3 position because the anticipated byproduct, ethyl butyrate, can be easily

removed.28 Benefits of this procedure include use of ambient temperature, neutral pH, and

conservative reaction time. The method is simple and “green”, as the lipase can be recycled.

Nevertheless, a significant amount of ethyl arachidonate is formed, due to over-hydrolysis. Since

the reaction is selective and proceeded quickly, it has also become a valuable tool for the

radiolabelled synthesis of 2-AG.29 The following work describes the extension of our method to

the synthesis of 2-acylglycerols, starting from saturated and unsaturated fatty acids, and alkyl

and aryl carboxylic acids.

3.2 Synthesis of 2-MAGs with Immobilized Candida antarctica and Rhizomucor miehei

3.2.1 Synthesis of 2-MAGs with Immobilized Candida antarctica

To test the general practicality of this lipase mediated scheme (Scheme 3.6), we

synthesized 2-MAGs from various commercially available long-chain carboxylic acids,

including those of biological importance. The synthesis began with the enzymatic 1,3-diacyl

protection of glycerol by the addition of CAL to glycerol and vinyl butyrate in anhydrous

CH2Cl2 at 0 °C. This resulted in the protected glycerol in quantitative yield.27, 30 The 1,3-

dibutyrylglycerol was then coupled to various medium and long-chain acids through EDCI

coupling in anhydrous CH2Cl2 with a catalytic amount of DMAP for four hours at 0 °C. This

generated the structured triglycerides (“ABA” type) in 67-99% yields.

For the hydrolysis step, CAL was added to the triglyceride, stirred in a minimal amount

of anhydrous ethanol at room temperature. By TLC analysis, it was observed that within one

hour the triglyceride had been completely consumed, and a mixture of 2-MAG, mono-protected

2-MAG (diacylglycerol), and ethyl butyrate was generated. There was no formation of ethyl

ester of fatty acid observed during this period. At this point, additional lipase was added to the

99

mixture, which was allowed to stir until all the diacylglycerol was consumed (1h), affording the

2-MAG. A significant amount of ethyl ester was observed during this period, and the formation

of ethyl ester largely depended on the type of fatty acid at the sn-2 position. Aryl and

unsaturated carboxylic acids showed more resistance towards over hydrolysis as compared to

saturated fatty acids.

Scheme 3.6 Chemoenzymatic Syntheses with immobilized Candida antarctica and Rhizomucor miehei

HO

OH

OH

Candida antarctica

99%

OCOC3H7

OCOC3H7

O

O 1EDCI, DMAP, CH2Cl2

R OH

O

56-99%

R O

OOCOC3H7

OCOC3H7

lipaseethanol

36-88%

R O

OOH

OH R O

OOH

OCOC3H7

1h

2a-14a

2b-14b

CH2Cl2 HO

R OEt

O

byproduct

Candida antarctica

Rhizomucor miehei

1h -24h

The separation of 1-MAG and 2-MAG is generally performed on boric acid impregnated

TLC plates and silica gel columns.31 Nonimpregnated silica TLC plates do not resolve 1- and 2-

MAGs. This separation is a necessary step for most 2-MAG syntheses due to the unfavorable

synthetic conditions used, where considerable formation of 1-MAG is observed. In contrast, the

highly regiospecific and neutral reaction conditions when using the lipase results in minimal or

no 1-MAG formation. Although silica gel purification has been reported to be a cause of acyl

migration from 2-MAG to 1-MAG,32 no migration was observed during column chromatography

with untreated silica gel. The only required step prior to purification was equilibration of silica

gel with hexanes. During chromatography, the highly non-polar ethyl ester byproduct eluted

100

with ethyl butyrate, and the 2-MAG was isolated without any acyl migration. It was observed

that the lipase-catalyzed hydrolysis reactions involving saturated triglycerides had isolated yields

<50%, with the ethyl ester byproduct being the major product, whereas the unsaturated

triglycerides had yields in the range of 55-75%, and triglycerides containing phenylalkyl groups

had yields >80% (Table 3.1). It should also be noted that there was no observable difference in

rate of reaction or isolated yield from the hydrolysis of a 1,3-diacetylglycerol-protected

compound as compared to the 1,3-dibutrylglycerol-protected compound.

3.2.2 Synthesis of 2-MAGs with Rhizomucor miehei

We screened a second 1,3-specific lipases to investigate whether the transformation can

be completed in better yields with a variety of substrates. Lipase from Rhizomucor miehei

showed excellent selectivity towards hydrolyzing “ABA” type triglycerides. The reaction

proceeded in a similar fashion where the triglyceride was consumed quickly, but hydrolysis of

the diglycerides took 24 to 48 hours. Even though the reaction proceeded very slowly compared

to CAL, the R. miehei lipase offered a remarkable improvement in selectivity, providing

exclusively 2-MAGs in excellent yields without formation of any ethyl ester byproduct from sn-

2 hydrolysis. The saturated and unsaturated fatty acid triglycerides were hydrolyzed in good

yields (75-88%) after 24 h. However, in most of the reactions, some unreacted diglyceride

intermediate remained. Allowing the reaction to proceed for an additional 24 h, or adding

additional lipase, did not increase the yields. When the unreacted diglyceride was separated

from the 2-MAG after 24 h, and was subjected to an additional treatment of Rhizomucor miehei

lipase the maximal yield once again reached `80% 2-MAG formation. Surprisingly, as in contrast

to CAL, Rhizomucor miehei lipase showed less reactivity towards aryl esters (13 and 14).

101

Table 3.1 Structure and Yields of Lipase catalyzed 2-MAGs

Cmpd. No. Triglyceride (a) 2-MAG (b)

C. antarctica

R. miehei

Yielda

(%) Yieldb

(%)

2 O

O

OCOC3H7

OCOC3H7H3C(H2C)10 O

O

OH

OHH3C(H2C)10

47 84

3 O

O

OCOC3H7

OCOC3H7H3C(H2C)12 O

O

OH

OHH3C(H2C)12

49 82

4 O

O

OCOC3H7

OCOC3H7H3C(H2C)14 O

O

OH

OHH3C(H2C)14

36 80

5 O

O

OCOC3H7

OCOC3H7

O

O

OH

OH

75 83

6 O

O

OCO(CH2)2CH3

OCO(CH2)2CH3H3C(H2C)16 O

O

OH

OHH3C(H2C)16

44 78

7 O

O

OCOC3H7

OCOC3H7

O

O

OH

OH

72 83

8 O

O

OCOC3H7

OCOC3H7

O

O

OH

OH

63 77

9 O

O

OCOC3H7

OCOC3H7

O

O

OH

OH

55 79

10 O

O

OCOC3H7

OCOC3H7

O

O

OH

OH

67 75

11 O

O

OCOCH3

OCOCH3

O

O

OH

OH

63 76

12 O

O

OCOC3H7

OCOC3H7H3C(H2C)20 O

O

OH

OHH3C(H2C)20

40 88

13 O

O

OCOC3H7

OCOC3H7

O

O

OH

OH

83 40

14 O

O

OCOC3H7

OCOC3H7

O

O

OH

OH

83 37

a Remaining yield consisted of the ethyl ester of sn-2 acyl group. b Remaining yield consisted of intermediate diglyceride.

102

3.3 Enzymatic synthesis of N-acylethanolamines

NAEs, ethanolamides of various long-chain fatty acids, constitute a class of bioactive

lipid molecules formed from glycerophospholipids through the phosphodiesterase-transacylation

pathway consisting of Ca2+-dependent N-acyltransferase and N-acylphosphatidylethanolamine-

hydrolyzing phospholipase D.33, 34 Among the NAEs, anandamide (Figure 3.1) is a

physiologically important lipid signaling molecule acting as a receptor ligand in the

endocannabinoid system.34 Recently, other NAEs such as PEA, and OEA (Figure 3.1) also

gained much attention due to their anti-inflammatory and analgesic activities, and anorexic

activity, respectively.35

O

NH

OH

O

NH

O

NH

H3C(H2C)14OH

OH

arachidonoylethanolamine(anandamide)

N-palmitoylethanolamine

N-oleoylethanolamine

Figure 3.1 N-Acylethanolamines

NAEs, including anandamide, are not stored in the cell but rather produced on demand,

and their endogenous levels are regulated directly by enzymes responsible for their formation

and degradation. Anandamide has a relatively rapid onset of action, but is also rapidly

hydrolyzed by FAAH, which accounts for its short duration of action. Early studies on structure-

activity relationships (SAR) focused on the preparation of various amides of arachidonic acid

and established that amides from chloroethylamine, cylopropylamine and R-(2)-aminopropanol

103

showed excellent improvement in their respective affinities to the cannabinoid CB1 receptor,

while exhibiting enhanced metabolic stability towards FAAH.36-40 Recently, SAR studies on the

modification of the hydrophobic chain have gained more attention and various analogs with fully

saturated fatty acid chains or alternatively encompassed alkyne moieties were synthesized.

Furthermore, our laboratory designed and synthesized high affinity covalent anandamide probes

for the CB1 receptor by introducing either electrophilic isothiocyanato or a photoactivatable

azido groups at the terminal carbon of the arachidonic acid moiety (Chapter 2).41 We also studied

the effect of aryl substitutions with variable methylene linker at the distal end of arachidonic

acid.42

All the synthetic schemes used the esters of substituted fatty acids as a starting point and

converted them to the needed amides using base mediated hydrolysis of the ester to carboxylic

acid, followed by activation of carboxylic acid with EDCI and treatment with various amines to

provide the respective amides. In a few cases, a protected form of ethanolamine was also used,

which required an additional deprotection step.

Several methods have been reported for the direct conversion of esters to amides

including magnesium methoxide,43 sodium cyanide,44 sodium methoxide,45 metal catalysts,46-48

and Alcalase.49 However, most of these methods suffer from incomplete conversion, longer

reaction times, and instability of the final products under the conditions used. Herein, we report a

highly selective lipase mediated mild conversion of esters to biologically important amides.

3.3.1 Candida antarctica for the direct aminolysis of esters

Lipases have found wide use as biocatalysts for many chemical transformations. Many

lipases have been studied for their use in amide formation,50-51 such as, amidation of benzyl

104

esters,52 synthesis of acetamides in the presence of ionic liquids,53 and acylation of amines with

acids.54 Most of these methods utilize either carboxylic acids or vinyl esters of carboxylic acids

as reactants and the reactions require relatively high temperatures. In the kinetic resolution of

amines, Nechab, et al., reported that the reaction conditions required 80 °C and 3-10 h to acylate

chiral amines with CAL and ethyl acetate.55 The aminolysis of linoleyl ethyl ester with

ethanolamine, catalyzed by CAL, in a solvent free system produced the linoleylethanolamide

only in 24% yield in 20 h, including the presence of the unwanted o-acylation product.56 While

these examples show the use of lipases for the amidation of esters, there is limited work reported

on the use of lipases as a direct method for the synthesis of biologically active NAEs with regard

to functional group sensitivity common in the synthesis of modified fatty acid moieties. We have

therefore focused our efforts on the synthesis of biologically active NAEs using immobilized

CAL from methyl esters and various amines. Developments in this area will imrpove the

synthesis of multistep tail-modified NAEs as well as other biologically important fatty acid

amide analogs.

3.3.2 Reaction optimization and results

To optimize the reaction conditions, we chose methyl arachidonate and cyclopropyl

amine as reactants and hexane as a solvent. When carried out at room temperature in the

presence of CAL the reaction proceeded smoothly, but very slowly, requiring 24 hrs for

completion. However, at 45 °C the reaction proceeded to completion significantly faster (3 hrs).

For amines that were not sufficiently soluble in n-hexane, the reaction proceeded equally well in

a 1:1 hexanes-diisopropyl ether mixture (Scheme 3.7). The results are shown in Table 3.2.

105

Scheme 3.7 CAL catalyzed aminolysis of esters

O

O

Immobilized Candida antartica

H2NOH

O

NH

OH

hexanes/diisopropylether 15

Table 3.2: Amidation of esters with immobilized Candida antarctica in 1:1 hexane-

diisopropylether

Amide

Entry Ester Amine Isolated yield Time

15

O

O

H2NOH

89% 2h

16

O

O

H2NOH

98% 24h

17

O

O

H2N 85% 3h

18

OH

O

O

H2NOH

85% 24h

19

N3

O

O

H2N 60% 24h

20 O

O

H2N

41% 24h

21 O

O

H2N 85% 3h

106

22 O

OH3C(H2C)14 H2N

84% 3h

23 O

O H2N 95% 24h

24 O

O

H2NOH

90% 24h

25

CO2Me

H2N

91% 24h

Esters and amines were chosen based on their biological importance. Methyl

arachidonate was treated with cyclopropyl amine, ethanolamine and (R)-2-aminopropanol to

provide ACPA, anandamide, and (R)-methanandamide, respectively, in excellent yields.

Unprotected ethanolamine was directly used in the preparation of various ethanolamides (2, 3, 4

and 10). When performed with a substituted fatty acid carrying a terminal hydroxyl group (4),

the reaction proceeded smoothly to provide the desired amide. There was no observable

transesterification product in any of the reactions where hydroxyl groups were present either in

the amine or the fatty acid substrates. In order to investigate the general applicability of the

method, we chose various esters and amines and showed that reactions proceeded smoothly with

esters. The yields were mainly dependant on the amine. Primary amines, including benzylic

amines, underwent amidation smoothly and in excellent yields. Conversely, cyclohexylamine

exhibited slower reactivity under the present conditions and longer reaction times and increased

temperature did not improve the yield significantly. Esters of non-fatty acids (9,10 and 11) also

107

underwent amidaton with amines in excellent yields and in all cases the reaction time appeared

to be more dependent on the amine used.

3.4 Conclusions

In summary, we have demonstrated that C. antarctica and R. miehei are suitable lipases

for the selective 1,3-hydrolysis of either an ‘AAA’ or ‘ABA’ triglyceride in order to produce

various 2-MAG. C. antarctica catalyzed reactions proceed faster, however, depending on the

substrates the yields were lower due to an ethyl ester by product. Meanwhile, R. miehei had

overall greater yields but required extended reaction times. In general, both are acceptable for

the chemoenzymatic synthesis of 2-MAGs. The work reported here has been recently

published.28, 57

CAL was also useful for the direct formation of amides from various amines and esters

containing skipped polyenes, allyl alcohol, allyl azide, alkyne, and aryl moieties. The method

described here, is simple, efficient and environmentally friendly, and does not require any

protection of other susceptible functional groups. This transacylation reaction provides excellent

yields and is selective. It may find general utility in the synthesis of amides from the

corresponding esters without requiring prior hydrolysis of the esters, as it can be difficult to

synthesize amides directly from esters under mild conditions. This method may prove useful in

the synthesis of drug intermediates and biologically important natural products. This work has

been recently published.58

108

3.5 Experimental

General. Statements from section 2.12 apply to the following experiments.

Compounds 3a-14a were synthesized following the procedure described for 2a; while

compounds 3b-14b were synthesized following the C. antarctica and R. miehei procedures

described for 2b.

OCOC3H7

OCOC3H7HO

2-hydroxypropane-1,3-diyl dibutyrate (1). Immobilized Candida antarctica (750 mg) was

added to a solution of glycerol (2.0 g, 21.6 mmol) and vinyl butyrate (6.2 g, 54.0 mmol) in

anhydrous CH2Cl2 (10 mL) at 0°C. The resulting mixture was stirred for 3h under argon

atmosphere. Then, additional lipase (400 mg) was added to the reaction mixture which was

stirred for an additional 2 h at 0°C. The lipase was filtered off, the solvent was evaporated off

under reduced pressure, and the residue was chromatographed on silica to yield 1 (5.0 g, 99%) as

a colorless oil. Rf = 0.55 (40% ethyl acetate/hexanes). 1H NMR (500 MHz, CHLOROFORM-d)

δ ppm 4.20 (dd, J=11.72, 4.39 Hz, 2 H) 4.14 (dd, J=11.72, 5.86 Hz, 2 H) 4.04 - 4.12 (m, 1 H)

2.41 - 2.58 (m, 1 H) 2.33 (t, J=7.32 Hz, 4 H) 1.67 (sxt, J=7.32 Hz, 4 H) 0.96 (t, J=7.57 Hz, 4 H).

The 13C NMR spectral data (100 MHz, CDCl3) are in agreement with literature values.27

O

O

OCO(CH2)2CH3

OCO(CH2)2CH3H3C(H2C)10

2-(dodecanoyloxy)propane-1,3-diyl dibutyrate (2a). EDCI (383 mg, 2.0 mmol), DMAP (19

mg, 0.16 mmol), and 1 (204 mg, 0.88 mmol) were added to a solution of lauric acid (160 mg,

0.80 mmol) in a 1:1 mixture of anhydrous THF/CH2Cl2 (10 mL) at 0 °C. The reaction was

109

allowed to stir for 4h. The reaction was then diluted with CH2Cl2 (15 mL) and H2O (15 mL).

The organic layer was separated, dried over MgSO4, and the solvent was removed under reduced

pressure. The residue was chromatographed on silica gel (0% to 15% ethyl acetate/hexanes) to

yield 2a (221 mg, 67%) as a colorless oil. Rf = 0.50 (15% ethyl acetate/hexanes). 1H NMR (400

MHz, CHLOROFORM-d) δ = 5.24 - 5.31 (m, 1 H) 4.30 (dd, J=12.09, 4.03 Hz, 2 H) 4.16 (dd,

J=12.46, 5.86 Hz, 2 H) 2.27 - 2.35 (m, 6 H) 1.57 - 1.70 (m, 6 H) 1.21 - 1.36 (m, 16 H) 0.92 -

0.98 (m, 6 H) 0.88 (t, J=6.60 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 173.4 (2C),

173.2, 77.4, 69.1, 62.3 (2C), 36.1 (2C), 34.4, 32.1, 29.8, 29.7, 29.6, 29.5, 29.3, 25.1, 22.9, 18.6

(2C), 14.4, 13.9 (2C). IR (neat) cm-1 2926, 2855, 1742, 1460. HRMS for C23H42O6Na (MNa+)

437.2881. Calcd. 437.2879.

O

O

OH

OHH3C(H2C)10

1,3-dihydroxypropan-2-yl dodecanoate (2b, utilizing Candida antarctica). Immobilized

Candida antarctica (Novozym 435, 100 mg) was added to a solution of 2a (100 mg, 0.24 mmol)

stirred in anhydrous EtOH (1 mL). After the full consumption of 2a (1h, TLC monitoring),

additional lipase (100 mg) was added until reaction completion was observed (1h). The reaction

mixture was diluted with CH2Cl2 (3 mL), and the lipase was filtered off. The solvent was

removed under reduced pressure, and the resulting residue was chromatographed on silica gel

(10% to 50% acetone/hexanes) to yield 2b (31mg, 47%) as a white solid. Rf = 0.26 (30%

acetone/hexanes). MP = 56-57 °C. 1H NMR (400 MHz, CHLOROFORM-d) δ = 4.93 (quin,

J=4.76 Hz, 1 H) 3.84 (br. s., 4 H) 2.38 (t, J=7.69 Hz, 2 H) 2.08 (br. s., 2 H) 1.58 - 1.69 (m, 2 H)

1.20 - 1.37 (m, 16 H) 0.88 (t, J=6.60 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ =

110

174.3, 75.3, 62.8 (2C), 34.6, 32.1, 29.8, 29.7, 29.6, 29.5, 29.3, 25.2 (2C), 22.9, 14.4. IR (neat)

cm-1 3352, 2922, 2856, 1730, 1464. HRMS for C15H30O4Na (MNa+) 297.2041. Calcd. 297.2042.

1,3-dihydroxypropan-2-yl dodecanoate (2b, utilizing Rhizomucor miehei). Lipozyme®,

immobilized from Rhizomucor miehei (100mg) was added to a solution of 2a (100 mg, 0.24

mmol) stirred in anhydrous EtOH (1 mL). The reaction was stirred for 24h, diluted with CH2Cl2

(3 mL), and the lipase was filtered off. The solvent was removed under reduced pressure, and

the resulting residue was chromatographed on silica gel (10% to 50% acetone/hexanes) to yield

2b (55 mg, 84%) as an oil. All spectral data was consistent with that obtained using the

procedure with C. antarctica.

O

O

OCO(CH2)2CH3


2-(tetradecanoyloxy)propane-1,3-diyl dibutyrate (3a). 347 mg, 99%, colorless oil. Rf = 0.47

(15% ethyl acetate/hexanes). 1H NMR (399MHz ,CHLOROFORM-d) δ = 5.32 - 5.22 (m, 1 H),

4.29 (dd, J = 4.4, 11.7 Hz, 2 H), 4.15 (dd, J = 5.9, 11.7 Hz, 2 H), 2.35 - 2.25 (m, 6 H), 1.70 - 1.56

(m, 6 H), 1.36 - 1.19 (m, 20 H), 0.94 (t, J = 7.3 Hz, 6 H), 0.87 (t, J = 6.6 Hz, 3 H). 13C NMR

(100MHz ,CHLOROFORM-d) δ = 173.4 (2C), 173.2, 69.1, 62.3 (2C), 36.1 (2C), 34.4, 32.2,

29.91, 29.89 (2C), 29.86, 29.7, 29.6, 29.5, 29.3, 25.1, 22.9, 18.6 (2C), 14.4, 13.9 (2C). IR (neat)

cm-1 2925, 2854, 1741, 1460. HRMS for C25H46O6Na (MNa+) 465.3195. Calcd. 465.3192.

O

O

OH

OHH3C(H2C)12

1,3-dihydroxypropan-2-yl tetradecanoate (3b). C. antarctica: 34mg, 49%; R. miehei: 57 mg,

82%; white solid. Rf = 0.22 (30% acetone/hexanes). MP = 57-58 °C. 1H NMR (399MHz

,CHLOROFORM-d) δ = 4.93 (quin, J = 4.8 Hz, 1 H), 3.89 - 3.78 (m, 3 H), 2.38 (t, J = 7.3 Hz, 2

111

H), 2.17 - 2.10 (m, 2 H), 1.70 - 1.58 (m, 2 H), 1.38 - 1.19 (m, 20 H), 0.88 (t, J = 7.3 Hz, 3 H). 13C

NMR (100MHz ,CHLOROFORM-d) δ = 174.3, 75.2, 62.8 (2C), 34.6, 32.2, 29.91, 29.87 (2C),

29.8, 29.7, 29.6, 29.5, 29.3, 25.2, 22.9, 14.4. IR (neat) cm-1 3418, 2926, 2855, 1729, 1466.

HRMS for C17H34O4Na (MNa+) 325.2354. Calcd. 325.2355.

O

O

OCO(CH2)2CH3


2-(palmitoyloxy)propane-1,3-diyl dibutyrate (4a). 309 mg, 84%, colorless oil. Rf = 0.39 (15%

ethyl acetate/hexanes). 1H NMR (500MHz ,CHLOROFORM-d) δ = 5.30 - 5.25 (m, 1 H), 4.30

(dd, J = 4.2, 12.0 Hz, 2 H), 4.16 (dd, J = 5.9, 11.7 Hz, 2 H), 2.35 - 2.27 (m, 6 H), 1.70 - 1.58 (m,

6 H), 1.34 - 1.21 (m, 24 H), 0.98 - 0.92 (m, 6 H), 0.88 (t, J = 6.8 Hz, 3 H). 13C NMR (100MHz

,CHLOROFORM-d) δ = 173.4, 173.2 (2C), 69.1, 62.3 (2C), 36.1, 34.4, 32.2, 29.9 (6C), 29.7,

29.6, 29.5, 29.3, 25.1 (2C), 18.6 (3C), 14.4, 13.9 (2C). IR (neat) cm-1 2924, 1742, 1460. HRMS

for C27H50O6Na (MNa+) 493.3503. Calcd. 493.3505.

O

O

OH

OHH3C(H2C)14

1,3-dihydroxypropan-2-yl palmitate (4b). C. antarctica: 25 mg, 36%; R. miehei: 56 mg, 80%;

white solid. Rf = 0.27 (30% acetone/hexanes). MP = 64-65 °C. 1H NMR (500MHz

,CHLOROFORM-d) δ = 4.93 (quin, J = 4.8 Hz, 1 H), 3.84 (t, J = 4.9 Hz, 4 H), 2.38 (t, J = 7.6

Hz, 2 H), 2.13 - 2.05 (m, 2 H), 1.69 - 1.59 (m, 2 H), 1.38 - 1.20 (m, 24 H), 0.88 (t, J = 7.3 Hz, 3

H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 174.3, 75.2, 62.8 (2C), 34.6, 32.2, 29.93,

29.92, 29.89, 29.84, 29.7, 29.6, 29.5, 29.3, 25.2 (2C), 23.3, 22.9, 14.4. IR (neat) cm-1 3320, 2917,

2850, 1730, 1471. HRMS for C19H38O4Na (MNa+) 353.2668. Calcd. 353.2668.

112

O

O

OCO(CH2)2CH3

OCO(CH2)2CH3

(Z)-2-(hexadec-9-enoyloxy)propane-1,3-diyl dibutyrate (5a). 291 mg, 79%, colorless oil. Rf =

0.50 (15% ethyl acetate/hexanes). 1H NMR (399MHz ,CHLOROFORM-d) δ = 5.38 - 5.32 (m, 2

H), 5.31 - 5.24 (m, 1 H), 4.30 (dd, J = 4.4, 11.7 Hz, 2 H), 4.16 (dd, J = 5.9, 12.5 Hz, 2 H), 2.38 -

2.26 (m, 6 H), 2.01 (q, J = 6.6 Hz, 4 H), 1.72 - 1.56 (m, 6 H), 1.39 - 1.20 (m, 16 H), 0.95 (t, J =

7.3 Hz, 6 H), 0.88 (t, J = 7.0 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 173.3 (2C),

173.1, 130.2, 129.9, 69.1, 62.3 (2C), 36.1 (2C), 34.4, 32.0, 30.0, 29.9, 29.4, 29.3, 29.25, 29.21,

27.5, 27.4, 25.1, 22.9, 18.6 (2C), 14.3, 13.8 (2C). IR (neat) cm-1 3007, 2928, 2856, 1742, 1459.


O

O

OH

OH

(Z)-1,3-dihydroxypropan-2-yl hexadec-9-enoate (5b). C. antarctica: 46 mg, 66%; R. miehei:

58 mg, 83%; colorless oil. Rf = 0.25 (30% acetone/hexanes). 1H NMR (399MHz

,CHLOROFORM-d) δ = 5.40 - 5.31 (m, 2 H), 4.92 (quin, J = 4.8 Hz, 1 H), 3.86 - 3.79 (m, 4 H),

2.37 (t, J = 7.7 Hz, 2 H), 2.33 (br. s., 2 H), 2.05 - 1.97 (m, 4 H), 1.63 (quin, J = 7.3 Hz, 2 H),

1.39 - 1.23 (m, 16 H), 0.88 (t, J = 6.6 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ =

174.3, 130.3, 129.9, 75.2, 62.6 (2C), 34.6, 32.0, 30.0, 29.9, 29.4, 29.32, 29.30, 29.2, 27.4, 27.4,

25.2, 22.9, 14.3. IR (neat) cm-1 3405, 3008, 2924, 2855, 1736, 1462. HRMS for C19H36O4Na

(MNa+) 351.2512. Calcd. 351.2511.

O

O

OCO(CH2)2CH3


113

2-(stearoyloxy)propane-1,3-diyl dibutyrate (6a). 300 mg, 85%, colorless oil. Rf = 0.34 (15%

ethyl acetate/hexanes). 1H NMR (500MHz ,CHLOROFORM-d) δ = 5.30 - 5.24 (m, 1 H), 4.30

(dd, J = 4.4, 11.7 Hz, 2 H), 4.16 (dd, J = 6.1, 12.0 Hz, 2 H), 2.34 - 2.27 (m, 6 H), 1.70 - 1.59 (m,

6 H), 1.35 - 1.20 (m, 28 H), 0.98 - 0.93 (m, 6 H), 0.88 (t, J = 6.6 Hz, 3 H). 13C NMR (100MHz

,CHLOROFORM-d) δ = 173.4 (2C), 173.2, 69.1, 62.3 (2C), 36.2 (2C), 34.4, 32.2, 29.9 (4C),

29.89 (3C), 29.86, 29.7, 29.6, 29.5, 29.3, 25.1, 22.9, 18.6 (2C), 14.4, 13.9 (2C). IR (neat) cm-1


O

O

OH

OHH3C(H2C)16

1,3-dihydroxypropan-2-yl stearate (6b). C. antarctica: 32mg, 44%; R. miehei: 56 mg, 78%;

white solid. Rf = 0.23 (30% acetone/hexanes). MP = 68-69 °C. 1H NMR (399MHz


H), 2.04 (t, J = 5.9 Hz, 2 H), 1.65 (quin, J = 7.3 Hz, 2 H), 1.38 - 1.19 (m, 28 H), 0.88 (t, J = 6.6

Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 174.3, 75.3, 62.8 (2C), 34.6, 32.2, 29.95

(5C), 29.91 (2C), 29.8, 29.7, 29.6, 29.5, 29.3, 25.2, 23.0, 13.8. IR (neat) cm-1 3313, 2916, 2849,

1730, 1472. HRMS for C21H42O4Na (MNa+) 381.2982. Calcd. 381.2981.

O

O

OCO(CH2)2CH3

OCO(CH2)2CH3

(Z)-2-(oleoyloxy)propane-1,3-diyl dibutyrate (7a). 290 mg, 82%, colorless oil. Rf = 0.43 (15%

ethyl acetate/hexanes). 1H NMR (500MHz ,CHLOROFORM-d) δ = 5.39 - 5.30 (m, 2 H), 5.30 -

5.24 (m, 1 H), 4.30 (dd, J = 4.4, 11.7 Hz, 2 H), 4.15 (dd, J = 6.1, 12.0 Hz, 2 H), 2.36 - 2.25 (m, 6

H), 2.01 (q, J = 6.2 Hz, 4 H), 1.71 - 1.55 (m, 6 H), 1.38 - 1.19 (m, 20 H), 0.95 (t, J = 7.3 Hz, 3

H), 0.88 (t, J = 6.8 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 173.4 (2C), 173.1,

114

130.3, 129.9, 69.1, 62.3, 36.1 (2C), 34.4, 32.1, 30.0, 29.9, 29.8, 29.6 (2C), 29.4, 29.3, 29.2, 27.5,

27.4, 25.1, 22.9, 18.6 (3C), 14.4, 13.9 (2C). IR (neat) cm-1 3007, 2925, 1742, 1460. HRMS for

C29H52O6Na (MNa+) 519.3658. Calcd. 519.3662.

O

O

OH

OH

1,3-dihydroxypropan-2-yl oleate (7b). C. antarctica: 48 mg, 67%; R. miehei: 60 mg, 83%;

colorless oil. Rf = 0.30 (30% acetone/hexanes). 1H NMR (399MHz ,CHLOROFORM-d) δ =

5.41 - 5.31 (m, 2 H), 4.92 (quin, J = 4.8 Hz, 1 H), 3.88 - 3.77 (m, 4 H), 2.49 (br. s., 2 H), 2.37 (t,

J = 7.7 Hz, 2 H), 2.01 (q, J = 6.4 Hz, 4 H), 1.63 (quin, J = 7.3 Hz, 2 H), 1.40 - 1.19 (m, 20 H),

0.88 (t, J = 6.6 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 174.4, 130.3, 129.9, 75.1,

62.5 (2C), 34.6, 32.1, 30.0, 29.9, 29.8, 29.6, 29.4, 29.33, 29.31, 27.45, 27.38, 25.2 (2C), 22.9,

14.4. IR (neat) cm-1 3415, 3008, 2923, 2854, 1735, 1464. HRMS for C21H40O4Na (MNa+)

379.2827. Calcd. 379.2824.

O

O

OCO(CH2)2CH3

OCO(CH2)2CH3

2-((9Z,12Z)-octadeca-9,12-dienoyloxy)propane-1,3-diyl dibutyrate (8a). 344 mg, 98%,

colorless oil. Rf = 0.38 (15% ethyl acetate/hexanes). 1H NMR (399MHz ,CHLOROFORM-d) δ =

5.43 - 5.30 (m, 4 H), 5.29 - 5.24 (m, 1 H), 4.30 (dd, J = 4.4, 11.7 Hz, 2 H), 4.15 (dd, J = 5.9, 11.7

Hz, 2 H), 2.77 (t, J = 6.6 Hz, 2 H), 2.36 - 2.26 (m, 6 H), 2.05 (q, J = 6.6 Hz, 4 H), 1.72 - 1.56 (m,

6 H), 1.41 - 1.22 (m, 14 H), 0.95 (t, J = 7.7 Hz, 6 H), 0.89 (t, J = 7.0 Hz, 3 H). 13C NMR

(100MHz ,CHLOROFORM-d) δ = 173.3 (2C), 173.1, 130.5, 130.2, 128.3, 128.1, 69.1, 62.3

(2C), 36.1 (2C), 34.4, 31.8, 29.8, 29.6, 29.4 (2C), 29.3, 29.2, 27.4, 25.8, 25.1, 22.8, 18.6 (2C),

115

14.3, 13.9 (2C). IR (neat) cm-1 3008, 2929, 2856, 1741, 1459. HRMS for C29H50O6Na (MNa+)

517.3506. Calcd. 517.3505.

O

O

OH

OH

(9Z,12Z)-1,3-dihydroxypropan-2-yl octadeca-9,12-dienoate (8b). C. antarctica: 45 mg, 63%;

R. miehei: 55 mg, 77%; colorless oil. Rf = 0.37 (30% acetone/hexanes). 1H NMR (399MHz


2.77 (t, J = 6.6 Hz, 2 H), 2.38 (t, J = 7.3 Hz, 2 H), 2.13 (t, J = 6.2 Hz, 2 H), 2.05 (q, J = 6.8 Hz, 4

H), 1.69 - 1.59 (m, 2 H), 1.41 - 1.23 (m, 14 H), 0.89 (t, J = 6.6 Hz, 3 H). 13C NMR (100MHz

,CHLOROFORM-d) δ = 174.3, 130.5, 130.2, 128.3, 128.1, 75.2, 62.8 (2C), 34.6, 31.8, 29.8,

29.6, 29.4, 29.33, 29.30, 27.4, 25.9, 25.2 (2C), 22.8, 14.3. IR (neat) cm-1 3397, 010, 2926, 2855,


O

O

OCO(CH2)2CH3

OCO(CH2)2CH3

(Z)-2-(icos-11-enoyloxy)propane-1,3-diyl dibutyrate (9a). 328 mg, 88%, colorless oil. Rf =


H), 5.30 - 5.24 (m, 1 H), 4.30 (dd, J = 4.4, 11.7 Hz, 2 H), 4.16 (dd, J = 5.9, 12.5 Hz, 2 H), 2.36 -

2.26 (m, 6 H), 2.05 - 1.97 (m, 4 H), 1.71 - 1.57 (m, 6 H), 1.27 (br. s., 24 H), 0.95 (t, J = 7.3 Hz, 6

H), 0.88 (t, J = 6.6 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 173.3 (2C), 173.1,

130.2, 130.0, 69.0, 62.3 (2C), 36.1 (2C), 34.4, 32.1, 30.0 (2C), 29.8, 29.7, 29.55 (2C), 29.52,

29.51, 29.3, 27.4 (2C), 25.1 (2C), 22.9, 18.6 (2C), 14.4, 13.9 (2C). IR (neat) cm-1 3008, 2925,


116

O

O

OH

OH

(Z)-1,3-dihydroxypropan-2-yl icos-11-enoate (9b). C. antarctica: 40mg, 55%; R. miehei, 58

mg, 79%; white solid. Rf = 0.24 (30% acetone/hexanes). MP = 32-33 °C. 1H NMR (399MHz


2.38 (t, J = 7.3 Hz, 2 H), 2.20 - 2.12 (m, 2 H), 2.01 (q, J = 6.6 Hz, 4 H), 1.71 - 1.57 (m, 2 H),

1.40 - 1.18 (m, 24 H), 0.88 (t, J = 6.6 Hz, 3 H). 13C NMR (100MHz ,CHLOROFORM-d) δ =

174.3, 130.2, 130.0, 75.2, 62.8 (2C), 34.6, 32.1, 30.0 (2C), 29.8, 29.7, 29.7, 29.6 (2C), 29.56,

29.51, 29.3, 27.4, 25.2 (2C), 23.0, 14.4. IR (neat) cm-1 3405, 3008, 2923, 2854, 1737, 1465.


O

O

OCO(CH2)2CH3

OCO(CH2)2CH3

2-((5Z,8Z,11Z,14Z)-icosa-5,8,11,14-tetraenoyloxy)propane-1,3-diyl dibutyrate (10a).

168mg, 98%, colorless oil. Rf = 0.36 (30% ethyl acetate/hexanes). The 1H and 13C spectral data

(500 and 100 MHz, CDCl3) are in agreement with literature values.28 IR (neat) 3012, 2931, 1741,

1456. HRMS for C31H50O6Na (MNa+) 541.3502. Calcd. 541.3505.

O

O

OH

OH

(5Z,8Z,11Z,14Z)-1,3-dihydroxypropan-2-yl icosa-5,8,11,14-tetraenoate (10b). C. antarctica:

48mg, 67%; R. miehei: 54 mg, 75%; colorless oil. Rf = 0.30 (30% acetone/hexanes). The 1H and

13C spectral data (500 and 100 MHz, CDCl3) are in agreement with literature values.28 IR (neat)

3420, 2013, 2927, 1736, 1456. HRMS for C23H38O4Na (MNa+) 401.2677. Calcd. 401.2668.

117

O

O

OCOCH3

OCOCH3

2-((4Z,7Z,10Z,13Z,16Z,19Z)-docosa-4,7,10,13,16,19-hexaenoyloxy)propane-1,3-diyl

diacetate (11a). EDCI (111 mg, 0.58 mmol), DMAP (6 mg, 0.06 mmol), and diacetin (44 mg,

0.25 mmol) were added to a solution of docosahexaenoic acid (75 mg, 0.23 mmol) in anhydrous

CH2Cl2 (5 mL) at 0 °C. The reaction was allowed to stir for 4h. Upon completion, the reaction

mixture was diluted with CH2Cl2 and H2O. The organic layer was separated, dried over MgSO4,

and removed under reduced pressure. The resulting residue was chromatographed on silica gel

(0% to 30% ethyl acetate/hexanes) to yield to 11a (111 mg, 99%) as a colorless oil. Rf = 0.55

(30% ethyl acetate/hexanes). 1H NMR (500MHz ,CHLOROFORM-d) δ = 5.47 - 5.32 (m, 12 H),

5.30 - 5.21 (m, 1 H), 4.29 (dd, J = 4.4, 11.7 Hz, 2 H), 4.16 (dd, J = 5.9, 12.2 Hz, 2 H), 2.93 - 2.77

(m, 10 H), 2.40 (d, J = 2.9 Hz, 4 H), 2.14 - 2.02 (m, 6 H), 0.98 (t, J = 7.6 Hz, 3 H). The 13C

spectral data (100 MHz, CDCl3) and IR data are in agreement with literature values.27 HRMS for

C29H42O6Na (MNa+) 509.2880. Calcd. 509.2879.

O

O

OH

OH

(4Z,7Z,10Z,13Z,16Z,19Z)-1,3-dihydroxypropan-2-yl docosa-4,7,10,13,16,19-hexaenoate

(11b). C. antarctica: 45 mg, 63%; R. miehei: 63 mg, 76%; colorless oil. Rf = 0.29 (30%

acetone/hexanes). 1H NMR (399MHz ,CHLOROFORM-d) δ = 5.52 - 5.22 (m, 12 H), 4.92 (quin,

J = 4.6 Hz, 1 H), 3.82 (t, J = 5.1 Hz, 4 H), 2.91 - 2.77 (m, 10 H), 2.49 - 2.37 (m, 4 H), 2.22 (t, J =

6.2 Hz, 2 H), 2.07 (quin, J = 7.5 Hz, 2 H), 0.97 (t, J = 7.3 Hz, 3 H). 13C NMR (100MHz

118

,CHLOROFORM-d) δ = 173.5, 132.3 (2C), 129.8 (2C), 128.8, 128.6, 128.5 (2C), 128.3 (2C),

128.1, 127.9, 75.4, 62.6 (2C), 34.4, 25.8 (5C), 23.0 (2C), 20.8. IR (neat) cm-1 3401, 3013, 2663,


O

O

OCO(CH2)2CH3


2-(docosanoyloxy)propane-1,3-diyl dibutyrate (12a). 201 mg, 56%, colorless oil. Rf = 0.48

(15% ethyl acetate/hexanes). MP = 27-28 °C. 1H NMR (399MHz ,CHLOROFORM-d) δ = 5.32 -

5.23 (m, 1 H), 4.30 (dd, J = 4.4, 11.7 Hz, 2 H), 4.16 (dd, J = 5.9, 11.7 Hz, 2 H), 2.36 - 2.26 (m, 6

H), 1.71 - 1.57 (m, 6 H), 1.25 (s, 36 H), 0.95 (t, J = 7.3 Hz, 6 H), 0.88 (t, J = 6.6 Hz, 3 H). 13C

NMR (100MHz ,CHLOROFORM-d) δ = 173.4 (2C), 173.2, 69.1, 62.3 (2C), 36.0 (2C), 34.4,

32.2, 29.95 (9C), 29.91 (2C), 29.88, 29.7, 29.6, 29.5, 29.3, 25.1, 22.9, 18.6 (2C), 14.4, 13.9 (2C).

IR (neat) cm-1 2923. 2853, 1742, 1462. HRMS for C33H62O6Na (MNa+) 577.4446. Calcd.

577.4444.

O

O

OCO(CH2)2CH3


1,3-dihydroxypropan-2-yl docosanoate (12b). C. antarctica: 32mg, 40%; R. miehei: 70 mg,

88%, white solid. Rf = 0.27 (30% acetone/hexanes). MP = 79-80 °C. 1H NMR (399MHz


H), 2.08 (s, 2 H), 1.69 - 1.59 (m, 2 H), 1.38 - 1.19 (m, 36 H), 0.88 (t, J = 6.2 Hz, 3 H). 13C NMR

(100MHz ,CHLOROFORM-d) δ = 174.3, 75.2, 62.8 (2C), 34.6, 32.2, 31.8, 29.94 (7C), 29.90

(2C), 29.8, 29.7, 29.6, 29.5, 29.3, 25.2, 22.94, 22.89, 14.4. IR (neat) cm-1 3313, 297, 2850, 1730,

1472. HRMS for C25H50O4Na (MNa+) 437.3610. Calcd. 437.3607.

119

O

O

OCO(CH2)2CH3

OCO(CH2)2CH3

2-(3-phenylpropanoyloxy)propane-1,3-diyl dibutyrate (13a). 280mg, 98%, colorless oil. Rf =


H), 7.24 - 7.15 (m, 3 H), 5.31 - 5.23 (m, 1 H), 4.28 (dd, J = 4.4, 11.7 Hz, 2 H), 4.13 (dd, J = 5.9,

11.7 Hz, 2 H), 2.96 (t, J = 7.7 Hz, 2 H), 2.66 (t, J = 8.1 Hz, 2 H), 2.34 - 2.24 (m, 6 H), 1.64 (sxt,

J = 7.3 Hz, 4 H), 0.94 (t, J = 7.3 Hz, 6 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 173.3

(2), 172.2, 140.4, 128.7 (2), 128.5 (2), 126.6, 69.4, 62.2 (2), 36.1 (2), 35.9, 31.0, 18.6 (2), 13.9

(2C). IR (neat) cm-1 3027, 2966, 2877, 1737, 1455. HRMS for C20H28O6Na (MNa+) 387.1787.

Calcd. 387.1784.

O

O

OH

OH

1,3-dihydroxypropan-2-yl 3-phenylpropanoate (13b). C. antarctica: 38mg, 83%; R. miehei: 8

mg, 40%; white foam. Rf = 0.18 (40% acetone/hexanes). 1H NMR (399MHz ,CHLOROFORM-

d) δ = 7.36 - 7.28 (m, 2 H), 7.26 - 7.18 (m, 3 H), 4.89 (td, J = 4.5, 9.3 Hz, 1 H), 3.79 - 3.71 (m, 4

H), 3.02 - 2.96 (m, 2 H), 2.77 - 2.70 (m, 2 H), 1.91 - 1.83 (m, 2 H). 13C NMR (100MHz

,CHLOROFORM-d) δ = 175.2, 140.4, 128.8 (2), 128.5 (2), 126.7, 75.5, 62.6 (2), 36.1, 31.8. IR

(neat) cm-1 3412, 3029, 2935, 2881, 1731, 1454. HRMS for C12H16O4Na (MNa+) 247.0945.

Calcd. 247.0946.

120

O

O

OCO(CH2)2CH3

OCO(CH2)2CH3

2-(5-phenylpentanoyloxy)propane-1,3-diyl dibutyrate (14a). 298mg, 99%, colorless oil. Rf =


H), 7.21 - 7.14 (m, 3 H), 5.31 - 5.23 (m, 1 H), 4.30 (dd, J = 4.4, 11.7 Hz, 2 H), 4.14 (dd, J = 5.9,

11.7 Hz, 2 H), 2.63 (t, J = 7.1 Hz, 2 H), 2.35 (t, J = 6.8 Hz, 2 H), 2.29 (t, J = 6.8 Hz, 4 H), 1.71 -

1.58 (m, 8 H), 0.94 (t, J = 7.3 Hz, 6 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 173.4 (2C),

172.9, 142.2, 128.6 (2C), 128.6 (2C), 126.0, 69.2, 62.3 (2C), 36.1 (2C), 35.8, 34.2, 31.0, 24.7,

18.6 (2C), 13.8 (2C). IR (neat) cm-1 3028, 2965, 2876, 1738, 1454. HRMS for C22H32O6Na

(MNa+) 415.2094. Calcd. 415.2097.

O

O

OH

OH

1,3-dihydroxypropan-2-yl 5-phenylpentanoate (14b). C. antarctica: 50mg, 83%; R. miehei:

24mg, 37%; white foam. Rf = 0.26 (40% acetone/hexanes). 1H NMR (399MHz

,CHLOROFORM-d) δ = 7.31 - 7.25 (m, 2 H), 7.22 - 7.13 (m, 3 H), 4.92 (td, J = 4.8, 9.5 Hz, 1

H), 3.86 - 3.76 (m, 4 H), 2.64 (t, J = 7.0 Hz, 2 H), 2.41 (t, J = 7.0 Hz, 2 H), 2.17 - 2.11 (m, 2 H),

1.74 - 1.60 (m, 4 H). 13C NMR (100MHz ,CHLOROFORM-d) δ = 174.0, 142.2, 128.6 (4C),

126.1, 75.2, 62.7 (2C), 35.8, 34.4, 31.0, 24.7. IR (neat) cm-1 3414, 3027, 2936, 2882, 1731, 1454.


121

O

NH

OH

(5Z,8Z,11Z,14Z)-N-(2-hydroxyethyl)icosa-5,8,11,14-tetraenamide (15).

Novozym 435 (100 mg) and ethanolamine (06 uL, 0.096 mmol) were added to a stirred solution

of methyl arachidonate (25 mg, 0.08 mmol) in a 1:1 mixture of hexanes and isopropyl ether (1

mL). The reaction was heated to 45 °C and stirred until completion (TLC monitoring, 2 h). The

reaction mixture was diluted with diethyl ether and filtered. The solvent was evacuated off under

reduced pressure and the resulting residue was chromatographed on silica gel to yield 15 (24 mg,

89%) as an oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 5.79 - 5.95 (m, 1H), 5.28 - 5.46 (m,

8H), 3.73 (q, J = 4.3 Hz, 2H), 3.41 - 3.45 (m, 2H), 2.78 - 2.87 (m, 6H), 2.22 (t, J = 7.3 Hz, 2H),

2.13 (q, J = 7.1 Hz, 2H), 2.06 (q, J = 7.3 Hz, 2H), 1.74 (quin, J = 7.5 Hz, 2H), 1.33 - 1.40 (m,

2H), 1.26 - 1.33 (m, 4H), 0.89 (t, J = 6.7 Hz, 3H).

O

NH

OH

(5Z,8Z,11Z,14Z)-N-((R)-1-hydroxypropan-2-yl)icosa-5,8,11,14-tetraenamide (16). The

procedure for 15 was followed to synthesize 16 (28 mg, 98%, 24 h). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 5.46 - 5.53 (m, 1 H) 5.29 - 5.44 (m, 8 H) 4.25 - 4.34 (m, 1 H) 4.13

(dd, J=11.0, 5.5 Hz, 1 H) 4.00 (dd, J=11.0, 4.3 Hz, 1 H) 2.73 - 2.90 (m, 7 H) 2.34 (t, J=7.6 Hz, 2

H) 2.15 (q, J=7.3 Hz, 2 H) 2.06 (q, J=6.7 Hz, 2 H) 1.71 (quin, J=7.5 Hz, 2 H) 1.36 (quin, J=6.7

Hz, 2 H) 1.25 - 1.32 (m, 4 H) 1.16 (d, J=6.7 Hz, 3 H) 0.89 (t, J=7.0 Hz, 3 H).

122

O

NH

(5Z,8Z,11Z,14Z)-N-cyclopropylicosa-5,8,11,14-tetraenamide (17). The procedure for 15 was

followed to synthesize 17 (mg, 85%, 3 h). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.56

(br. s., 1 H) 5.24 - 5.46 (m, 8 H) 2.82 (dq, J=12.4, 6.1 Hz, 6 H) 2.70 (tq, J=7.1, 3.6 Hz, 1 H) 2.09

- 2.15 (m, 4 H) 2.01 - 2.08 (m, 2 H) 1.70 (quin, J=7.5 Hz, 2 H) 1.33 - 1.40 (m, 2 H) 1.25 - 1.33

(m, 4 H) 0.89 (t, J=6.7 Hz, 2 H) 0.74 - 0.79 (m, 2 H) 0.45 - 0.50 (m, 2 H).

OH

O

NH

OH

(5Z,8Z,11Z)-13-hydroxy-N-(2-hydroxyethyl)trideca-5,8,11-trienamide (18). The procedure

for 15 was followed to synthesize 18 (mg, 85%, 24 h). 1H NMR (500 MHz, CHLOROFORM-d)

δ ppm 6.32 (br. s., 1 H) 5.58 - 5.71 (m, 1 H) 5.47 - 5.57 (m, 1 H) 5.25 - 5.46 (m, 4 H) 4.22 (d,

J=6.7 Hz, 2 H) 3.70 (t, J=4.9 Hz, 2 H) 3.40 (q, J=5.5 Hz, 2 H) 2.87 (t, J=6.4 Hz, 2 H) 2.81 (t,

J=6.1 Hz, 2 H) 2.22 (t, J=7.3 Hz, 2 H) 2.09 - 2.15 (m, 2 H) 1.72 (quin, J=7.5 Hz, 2 H).

N3

O

NH

(5Z,8Z,11Z)-13-azido-N-cyclopropyltrideca-5,8,11-trienamide (19). The procedure for 15

was followed to synthesize 19 (mg, 60%, 24 h). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm

5.66 - 5.86 (m, 1 H) 5.48 - 5.64 (m, 1 H) 5.33 - 5.46 (m, 4 H) 3.69 - 3.75 (m, 2 H) 2.86 (t, J=5.8

Hz, 2 H) 2.75 - 2.80 (m, 2 H) 2.70 (tq, J=7.0, 3.5 Hz, 1 H) 2.06 - 2.15 (m, 4 H) 1.70 (quin, J=7.5

Hz, 2 H) 0.74 - 0.79 (m, 2 H) 0.45 - 0.51 (m, 2 H).

123

O

NH

N-cyclohexyloleamide (20). The procedure for 15 was followed to synthesize 20 (mg, 41%, 24

h). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.29 - 5.39 (m, 2 H) 2.36 (br. s., 2 H) 1.98 -

2.04 (m, 4 H) 1.58 - 1.67 (m, 2 H) 1.22 - 1.37 (m, 24 H) 0.88 (t, J=7.3 Hz, 3 H).

O

NH

(4Z,7Z,10Z,13Z,16Z,19Z)-N-cyclopropyldocosa-4,7,10,13,16,19-hexaenamide (21). The

procedure for 15 was followed to synthesize 21 (mg, 85%, 3 h). 1H NMR (399 MHz,

CHLOROFORM-d) δ ppm 5.61 (br. s., 1 H) 5.23 - 5.47 (m, 12 H) 2.75 - 2.91 (m, 10 H) 2.70

(ddt, J=10.8, 7.1, 3.1, 3.1 Hz, 1 H) 2.40 (q, J=7.1 Hz, 2 H) 2.18 (t, J=7.3 Hz, 2 H) 2.02 - 2.12 (m,

2 H) 0.97 (t, J=6.6 Hz, 3 H) 0.73 - 0.80 (m, 2 H) 0.44 - 0.51 (m, 2 H).

O

NH

H3C(H2C)14

N-(prop-2-ynyl)palmitamide (22). The procedure for 15 was followed to synthesize 22 (mg,

84%, 3 h). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.51 - 5.60 (m, 1 H) 4.06 (dd, J=5.2,

2.7 Hz, 2 H) 2.23 (t, J=2.4 Hz, 1 H) 2.19 (t, J=7.3 Hz, 2 H) 1.59 - 1.67 (m, 2 H) 1.20 - 1.33 (m,

24 H) 0.88 (t, J=7.0 Hz, 3 H)

O

NH

N-benzylhex-5-ynamide (23). The procedure for 15 was followed to synthesize 23 (mg, 95%,

24 h). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.32 - 7.37 (m, 2 H) 7.27 - 7.31 (m, 3 H)

124

5.75 (br. s., 1 H) 4.45 (d, J=6.1 Hz, 2 H) 2.37 (t, J=7.3 Hz, 2 H) 2.28 (td, J=6.9, 2.7 Hz, 2 H)

1.96 (t, J=2.7 Hz, 1 H) 1.90 (quin, J=7.0 Hz, 2 H).

O

NH

OH

N-(2-hydroxyethyl)-3-phenylpropanamide (24). The procedure for 15 was followed to

synthesize 24 (mg, 90%, 24 h). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.27 - 7.33 (m, 2

H) 7.18 - 7.24 (m, 3 H) 5.79 (br. s., 1 H) 3.63 (q, J=4.9 Hz, 2 H) 3.33 - 3.38 (m, 2 H) 2.98 (t,

J=7.6 Hz, 2 H) 2.51 (t, J=7.6 Hz, 2 H) 2.40 (t, J=5.2 Hz, 1 H).

O

NH

N-benzylbenzamide (25). The procedure for 15 was followed to synthesize 25 (mg, 91%, 24 h).

1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.34 - 7.37 (m, 2 H) 7.27 - 7.32 (m, 5 H) 7.19 -

7.24 (m, 3 H) 5.82 (br. s., 1 H) 4.47 (d, J=6.1 Hz, 2 H).

125

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15. Berger, M.; Schnelder, M. Enzymatic esterification of glycerol II. Lipase-catalyzed synthesis of regioisomerically pure 1(3)-rac -monoacylglycerols. Journal of the American Oil Chemists' Society 1992, 69, 961-965. 16. Piyatheerawong, W.; Yamane, T.; Nakano, H.; Iwasaki, Y. Enzymatic preparation of enantiomerically pure sn -2,3-diacylglycerols: A stereoselective ethanolysis approach. Journal of the American Oil Chemists' Society 2006, 83, 603-607. 17. Irimescu, R.; Yasui, M.; Iwasaki, Y.; Shimidzu, N.; Yamane, T. Enzymatic synthesis of 1,3-dicapryloyl-2-eicosapentaenoylglycerol. Journal of the American Oil Chemists' Society 2000, 77, 501-506. 18. Rosu, R.; Yasui, M.; Iwasaki, Y.; Yamane, T. Enzymatic synthesis of symmetrical 1,3-diacylglycerols by direct esterification of glycerol in solvent-free system. Journal of the American Oil Chemists' Society 1999, 76, 839-843. 19. Schmid, U.; Bornscheuer, U. T.; Soumanou, M. M.; McNeill, G. P.; Schmid, R. D. Highly selective synthesis of 1,3-oleoyl-2-palmitoylglycerol by lipase catalysis. Biotechnology and Bioengineering 1999, 64, 678-684. 20. Byun, H.-G.; Eom, T.-K.; Jung, W.-K.; Kim, S.-K. Lipase catalyzed production of monoacylglycerols by the esterification of fish oil fatty acids with glycerol. Biotechnology and Bioprocess Engineering 2007, 12, 491-496. 21. Waldinger, C.; Schneider, M. Enzymatic esterification of glycerol III. Lipase-catalyzed synthesis of regioisomerically pure 1,3-sn -diacylglycerols and 1 (3)- rac -monoacylglycerols derived from unsaturated fatty acids. Journal of the American Oil Chemists' Society 1996, 73, 1513-1519. 22. Schmid, U.; Bornscheuer, U.; Soumanou, M.; McNeill, G.; Schmid, R. Optimization of the reaction conditions in the lipase-catalyzed synthesis of structured triglycerides. Journal of the American Oil Chemists' Society 1998, 75, 1527-1531. 23. Soumanou, M. M.; Bornscheuer, U. T.; Schmid, U.; Schmid, R. D. Crucial Role of Support and Water Activity on the Lipase-Catalyzed Synthesis of Structured Triglycerides. Biocatalysis and Biotransformation 1999, 16, 443-459. 24. Soumanou, M.; Bornscheuer, U.; Schmid, R. Two-step enzymatic reaction for the synthesis of pure structured triacylglycerides. Journal of the American Oil Chemists' Society 1998, 75, 703-710. 25. Wongsakul, S.; Prasertsan, P.; Bornscheuer, U. T.; H-Kittikun, A. Synthesis of 2-monoglycerides by alcoholysis of palm oil and tuna oil using immobilized lipases. European Journal of Lipid Science and Technology 2003, 105, 68-73. 26. Irimescu, R.; Iwasaki, Y.; Hou, C. Study of TAG ethanolysis to 2-MAG by immobilized Candida antarctica lipase and synthesis of symmetrically structured TAG. Journal of the American Oil Chemists' Society 2002, 79, 879-883. 27. Devane, W. A.; Dysarz, F. A.; Johnson, M. R.; Melvin, L. S.; Howlett, A. C. Determination and characterization of a cannabinoid receptor in rat brain. Molecular Pharmacology 1988, 34, 605-613. 28. Vadivel, S. K.; Whitten, K. M.; Makriyannis, A. Chemoenzymatic synthesis of 2-arachidonoylglycerol, an endogenous ligand for cannabinoid receptors. Tetrahedron Letters 2011, 52, 1149-1150. 29. Duclos, R. I.; Johnston, M.; Vadivel, S. K.; Makriyannis, A.; Glaser, S. T.; Gatley, S. J. A Methodology for Radiolabeling of the Endocannabinoid 2-Arachidonoylglycerol (2-AG). The Journal of Organic Chemistry 2011, 7, 2049–2055.

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30. Halldorsson, A.; Magnusson, C. D.; Haraldsson, G. G. Chemoenzymatic synthesis of structured triacylglycerols by highly regioselective acylation. Tetrahedron 2003, 59, 9101-9109. 31. Takagi, T.; Ando, Y. Stereospecific analysis of triacyl-<i>sn</i>-glycerols by chiral high-performance liquid chromatography. Lipids 1991, 26, 542-547. 32. Irimescu, R.; Furihata, K.; Hata, K.; Iwasaki, Y.; Yamane, T. Two-step enzymatic synthesis of docosahexaenoic acid-rich symmetrically structured triacylglycerols <i>via</i> 2-monoacylglycerols. Journal of the American Oil Chemists' Society 2001, 78, 743-748. 33. Coulon, D.; Faure, L.; Salmon, M.; Wattelet, V.; Bessoule, J.-J. N-Acylethanolamines and related compounds: Aspects of metabolism and functions. Plant Science 2012, 184, 129-140. 34. Ezzili, C.; Otrubova, K.; Boger, D. L. Fatty acid amide signaling molecules. Bioorganic & Medicinal Chemistry Letters 2010, 20, 5959-5968. 35. Ueda, N.; Tsuboi, K.; Uyama, T. Enzymological studies on the biosynthesis of N-acylethanolamines. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids 2010, 1801, 1274-1285. 36. Abadji, V.; Lin, S.; Taha, G.; Griffin, G.; Stevenson, L. A.; Pertwee, R. G.; Makriyannis, A. (R)-Methanandamide: A Chiral Novel Anandamide Possessing Higher Potency and Metabolic Stability. Journal of Medicinal Chemistry 1994, 37, 1889-1893. 37. Goutopoulos, A.; Fan, P.; Khanolkar, A. D.; Xie, X.-Q.; Lin, S.; Makriyannis, A. Stereochemical Selectivity of Methanandamides for the CB1 and CB2 Cannabinoid Receptors and Their Metabolic Stability. Bioorganic & Medicinal Chemistry 2001, 9, 1673-1684. 38. Bezuglov, V.; Bobrov, M.; Gretskaya, N.; Gonchar, A.; Zinchenko, G.; Melck, D.; Bisogno, T.; Di Marzo, V.; Kuklev, D.; Rossi, J.-C.; Vidal, J.-P.; Durand, T. Synthesis and biological evaluation of novel amides of polyunsaturated fatty acids with dopamine. Bioorganic & Medicinal Chemistry Letters 2001, 11, 447-449. 39. El Fangour, S.; Balas, L.; Rossi, J.-C.; Fedenyuk, A.; Gretskaya, N.; Bobrov, M.; Bezuglov, V.; Hillard, C. J.; Durand, T. Hemisynthesis and preliminary evaluation of novel endocannabinoid analogues. Bioorganic & Medicinal Chemistry Letters 2003, 13, 1977-1980. 40. Urbani, P.; Cavallo, P.; Cascio, M. G.; Buonerba, M.; De Martino, G.; Di Marzo, V.; Saturnino, C. New metabolically stable fatty acid amide ligands of cannabinoid receptors: Synthesis and receptor affinity studies. Bioorganic & Medicinal Chemistry Letters 2006, 16, 138-141. 41. Li, C.; Xu, W.; Vadivel, S. K.; Fan, P.; Makriyannis, A. High affinity electrophilic and photoactivatable covalent endocannabinoid probes for the CB1 receptor. J Med Chem 2005, 48, 6423-9. 42. Yao, F.; Li, C.; Vadivel, S. K.; Bowman, A. L.; Makriyannis, A. Development of novel tail-modified anandamide analogs. Bioorganic & Medicinal Chemistry Letters 2008, 18, 5912-5915. 43. Bundesmann, M. W.; Coffey, S. B.; Wright, S. W. Amidation of esters assisted by Mg(OCH3)2 or CaCl2. Tetrahedron Lett. 2010, 51, 3879-3882. 44. Hoegberg, T.; Stroem, P.; Ebner, M.; Raemsby, S. Cyanide as an efficient and mild catalyst in the aminolysis of esters. Journal of Organic Chemistry 1987, 52, 2033-2036. 45. Ohshima, T.; Hayashi, Y.; Agura, K.; Fujii, Y.; Yoshiyama, A.; Mashima, K. Sodium methoxide: a simple but highly efficient catalyst for the direct amidation of esters. Chemical Communications 2012, 48, 5434-5436.

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46. Gnanaprakasam, B.; Milstein, D. Synthesis of amides from esters and amines with liberation of H2 under neutral conditions. J. Am. Chem. Soc. 2011, 133, 1682-1685. 47. Han, C.; Lee, J. P.; Lobkovsky, E.; Porco, J. A. Catalytic Ester−Amide Exchange Using Group (IV) Metal Alkoxide−Activator Complexes. Journal of the American Chemical Society 2005, 127, 10039-10044. 48. Ishihara, K.; Kuroki, Y.; Hanaki, N.; Ohara, S.; Yamamoto, H. Antimony-Templated Macrolactamization of Tetraamino Esters. Facile Synthesis of Macrocyclic Spermine Alkaloids, (±)-Buchnerine, (±)-Verbacine, (±)-Verbaskine, and (±)-Verbascenine. Journal of the American Chemical Society 1996, 118, 1569-1570. 49. Nuijens, T.; Cusan, C.; Kruijtzer, J. A. W.; Rijkers, D. T. S.; Liskamp, R. M. J.; Quaedflieg, P. J. L. M. Enzymatic Synthesis of C-Terminal Arylamides of Amino Acids and Peptides. Journal of Organic Chemistry 2009, 74, 5145-5150. 50. Gotor, V. Non-conventional hydrolase chemistry: amide and carbamate bond formation catalyzed by lipases. Bioorganic & Medicinal Chemistry 1999, 7, 2189-2197. 51. Bistline, R.; Bilyk, A.; Feairheller, S. Lipase catalyzed formation of fatty amides. Journal of the American Oil Chemists' Society 1991, 68, 95-98. 52. Adamczyk, M.; Grote, J. Pseudomonas cepacia lipase mediated amidation of benzyl esters. Tetrahedron Letters 1996, 37, 7913-7916. 53. Dhake, K. P.; Qureshi, Z. S.; Singhal, R. S.; Bhanage, B. M. Candida antarctica lipase B-catalyzed synthesis of acetamides using [BMIm(PF6)] as a reaction medium. Tetrahedron Letters 2009, 50, 2811-2814. 54. Tufvesson, P.; Annerling, A.; Hatti-Kaul, R.; Adlercreutz, D. Solvent-free enzymatic synthesis of fatty alkanolamides. Biotechnology and Bioengineering 2007, 97, 447-53. 55. Nechab, M.; Azzi, N.; Vanthuyne, N.; Bertrand, M.; Gastaldi, S.; Gil, G. Highly selective enzymatic kinetic resolution of primary amines at 80 degrees C: a comparative study of carboxylic acids and their ethyl esters as acyl donors. Journal of Organic Chemistry 2007, 72, 6918-23. 56. Couturier, L.; Taupin, D.; Yvergnaux, F. Lipase-catalyzed chemoselective aminolysis of various aminoalcohols with fatty acids. Journal of Molecular Catalysis B: Enzymatic 2009, 56, 29-33. 57. Whitten, K. M.; Makriyannis, A.; Vadivel, S. K. Application of chemoenzymatic hydrolysis in the synthesis of 2-monoacylglycerols. Tetrahedron 2012, 68, 5422-5428. 58. Whitten, K. M.; Makriyannis, A.; Vadivel, S. K. Enzymatic synthesis of N-Acylethanolamines: Direct method for the aminolysis of esters. Tetrahedron Letters 2012, 53, 5753-5755.

129

CHAPTER 4

CHEMICALLY AND METABOLICALLY STABLE 2-ARACHIDONOYLGLYCEROL

ANALOGS

130

4.1 Introduction

Through the use of chemoenzymatic methods we minimized the acyl migration of 2-AG

during synthesis, however, the chemical and metabolic instability of 2-AG is an issue during

storage, in vitro assays, and during in vivo pharmacological profile experiments. The chemical

instability of 2-AG leads to acyl migration to 1-AG, which can produce different effects, while

metabolic instability results in rapid hydrolysis to arachidonic acid and glycerol. The first

reported stable 2-AG analog discovered was 2-arachidonyl glycerol ether1 (2-AGE, Figure 4.1);

which was synthesized and evaluated by Sugiura, et al.2 2-AGE is an analog with comparable

although weaker) cannabinergic in vitro and in vivo properties than 2-AG. This observation led

to the discussion of 2-AGE being labeled as the third endocannabinoid.3

A common technique to improve metabolic stability of a compound is the introduction of

steric moieties at the site of hydrolysis.4 This technique was tested with 2-AG by α-methylation

of the glycerol ester carbonyl (1, Figure 4.1).5 At the same time the glycerol hydroxyl groups

were replaced with fluorine in an attempt to prevent acyl migration, as seen with 2 and 3.

However, the introduction of this methyl resulted in a significant decrease in CB1 affinity as

compared to 2-AG. α-Methylation was also unsuccessful in preventing acyl migration in a

membrane free buffer solution, although a 50% decrease in hydrolysis to arachidonic acid was

observed. Mono-fluorination of 2-AG, 2, showed a dramatic decrease in affinity, while 3 had

even less activity at the CB receptor. Parkkari, et al., replaced the -C5H11 tail of 2-AG with a

dimethylheptyl group, however, no improvement in activity or efficacy over 2-AG was

observed.6 These results support the importance of the free hydroxyl groups of the glycerol as

crucial features for high affinity and efficacy of 2-AG type compounds in the CB system.

131

O

O

OH

OHO

O

OH

FO

O

F

F

O

OH

OH

OH

OH

OH

OH

OH

OHO OH

O

OHO

OHO

OHO

OHO

OHO

OH

meso-7 meso-9±8

2-AG

1 2 3

4 5 6

HO OO

O

OH

OH10

5 10

O

OH

OH

2-AGE

O

Figure 4.1 2-AG and published 2-AG analogs

The most obvious method to prevent hydrolysis would be to remove the ester, as seen in

2-AGE. Suhara, et al., did this by replacing the ester with a ketone, as in 4, followed by a

reduction of the ketone to the racemic alcohol (5), and finally the conversion to straight alkyl

linkage in 6. While this increased metabolic stability, it markedly reduced the binding affinity as

compared to 2-AG.7

132

Our lab has synthesized conformationally constrained 2-AG analogs designed to increase

chemical and metabolic stability. Meso-7 showed minimal hydrolysis by FAAH and MGL,

while 8 and meso-9 were only stable towards MGL. However, their binding affinities for the CB

receptors were reduced as compared to 2-AG.8

More recently, Brizzi, et al., developed resorcinol-sn-glycerol derivatives as 2-AG

analogs.9 Compound 10 displayed exceptional CB1 binding affinity (10 nM), however, this class

of compounds exhibited very low efficacy in multiple assays. The authors’ findings also

suggested that these compounds possessed acceptable metabolic stability. The weak functional

activity of the glycerol compounds was proposed to be from rapid acyl migration to the 1-AG

variant of the analog which would produce reduced activity.

The preparation of 2-AG analogs that maintain sufficient affinity and efficacy in vitro

and in vivo while simultaneously surmounting chemical and metabolic instabilities has so far

been unsuccessful. The identification of a stable 2-AG analog is critical for understanding the

pharmacological profile and complete biological function.

4.2 Synthesis of Biphenyl 2-AG Analogs

The first strategy was to synthesize 2-AG analogs that replaced the fatty acid backbone

with a biphenyl moiety. In endocannabinoids the unsaturated region of the arachidonic chain

folds itself into a ‘U’ or ‘J’ conformation10 in which the alkyl chain wraps itself around and the

C5H11 tail is in the vicinity of the ester head-group. It’s hypothesized that the length of the tail

influences the acyl migration of the glycerol ester of 2-MAG, as the rate of acyl migration has

observed to decrease with shorter fatty acid chains.11 The arachidonic chain is also believed to

mimic the tricyclic structure of THC,12 therefore, by replacing a long saturated chain with a

133

biphenyl moiety we may be able to reduce acyl migration while maintaining cannabinoid

potency and efficacy.

The first analogs were synthesized from biphenyl-4-carboxaldehyde 11. Treatment with

methyl (triphenylphosphoranylidene)acetate yielded the Wittig product 12 as mostly the trans

isomer (Scheme 4.1). While the cis isomer was separated from 12, ensuing hydrolysis

isomerized the Z-alkene to result in 13. Biphenyl acid 14 was the product of 11 treated with (2-

carboxyethyl)triphenylphosphonium bromide and potassium tert-butoxide.

Scheme 4.1 Synthesis of biphenyl 2-AG analogs

H

O

O

O

O

On

O

O

O

O

O

On

OH

O

O

O

On

OH

OH

O

O

O

O

O

O

O

O

O

O

OHO

O

OH

OH

O

OHn

13 n = 014 n = 1

Ph3 P

O

O

51%

Ph3P

O

OHBr

KOtBu, THF, -78 °C

46%

LiOH, THF57%

EDCI, 1,3-DBGDMAP, DCM

83-84%

15 n = 016 n = 1

C. antarcticaethanol

35-67%

17 n = 019 n = 1 18 n = 0

12Pd/C, H2, EtOAc

82%

O

O

O

OHLiOH, THF

63%

EDCI, 1,3-DBGDMAP, DCM

61%


59%/35%

20 21

22 23 24

11

12

In performing these Wittig reactions, use of an ylide or phosphonium salt containing an

acid, as opposed to the methyl ester, was preferred in order to bypass the hydrolysis step.

134

However, ylides of methyl esters were the only stable ylides available from commercial sources.

Otherwise, phosphonium bromides containing an acid were vigorously dried under vacuum with

phosphorous pentoxide prior to use.

Acids 13 and 14 were esterified with 1,3-dibutyrylglycerol (1,3-DBG) in the presence of EDCI

and DMAP to yield 15 and 16. Treatment of the ‘ABA’ triglycerides with CAL yielded

diglycerides 17 and 19 along with 2-MAG 18 when the reaction was stopped after 1 h. As

observed in Chapter 3, additional reaction time resulted in full consumption of the diglyceride.

Compound 12 was treated with palladium on carbon and hydrogen. And after hydrolysis,

esterification, and lipase induced-hydrolysis, diglyceride 23 and 2-MAG 24 were isolated.

While the Wittig method is useful for creating shorter biphenyl compounds, synthesizing

analogs with longer carbon chains can be cumbersome, as the hygroscopicity of ylides and

Wittig salts can reduce reaction yields. To continue this method the phosphonium bromide salts

would be synthesized and vigorously dried prior to use.

As an alternative to the Wittig method, a microwave-mediated Sonagashira reaction was

investigated. The terminal alkynes were commercially available and therefor preparation of

reagents is minimal. This method also allows for easy conversion of the alkyne to the cis-alkene

through partial hydrogenation.

The reaction of 4-bromobiphenyl 25 with methyl 5-hexynoate 26,

bis(triphenylphosphine)palladium(II) chloride, copper iodide, and triethylamine using

microwave irradiation gave ester 27 in good yield (Scheme 4.2). When this reaction was

attempted using the acid as opposed to the ester, the yields were markedly reduced. Therefore,

ester 27 was hydrolyzed with lithium hydroxide to give acid 28 which was then coupled to 1,3-

DBG utilizing carbodiimide conditions to yield ‘ABA’ triglyceride 29. C. antarctica was

135

applied in two separate reactions, with the first being stopped after 1 h to yield diglyceride 30 in

79% yield, and a second reaction allowed to go to completion to yield 2-MAG 31 in 78% yield.

Scheme 4.2 Continued synthesis of biphenyl 2-AG analogs

BrO

O

O

OH

O

O

O

O

O

O

O

O

O

O

OH

O

O

OH

OH

O

O

O

O

O

OO

O

O

O

O

O

O

O

O

O

OH

O

O

OH

OH

O

O

O

O

OH

O

O

OH

OH

25 27 28

29

30

31

32 33

34

35

36

37

Pd(PPh3)2Cl2Et3N, CuI, DMF

62%

LiOH, THF

91%

1,3-DBG, DCMEDCI, DMAP

80%


78-79%


8-65%

Pd/C, H2ether

76%


79%

O

O26

Triglyceride 29 was fully saturated to give 32, after which treatment with CAL yielded

the diglyceride and 2-MAG, 34 and 35. To provide the cis-alkene, 29 was treated with P-2 nickel

136

catalyst (created in situ) and hydrogen to yield 33 in 79% yield. This gave 36 and 37 after

reaction with CAL.

For the next analogs the goal was to insert an ether linkage between the core benzene

rings. This substitution would decrease the hydrophobicity of these 2-AG analogs, possibly

increasing the ability to permeate cells. 2-AG and anandamide are brought into cells by

transporters13 due to their high lipophilicity.

Scheme 4.3 Synthesis of biphenyl ether 2-AG analogs

OH

O 1,3-DBG, DCMEDCI, DMAP

72%

O

OO

O

O

O

O

O

O

O

O

O

O

O

Br Pd(PPh3)2Cl2Et3N, CuI, DMF

42%


O

O

O

OH

OH


63%

O

O

O

O

O

O

OC. antarcticaethanol O

O

OH

OH

O

79%

75%

38 39 40

41 42

43 44

As previous Sonagashira reactions with acid-containing alkynes were poor, 4-pentynoic

acid 38 was esterified with 1,3-DBG to yield 39 (Scheme 4.3). 39 was then subjected to

Sonagashira conditions using microwave irradiation with 4-bromobiphenyl ether 40 to yield

137

triglyceride 41. 41 was partially hydrogenated with P-2 nickel to yield cis-alkene 43. 41 and 43

were treated with CAL to yield 2-MAG 42 and 44 respectively in good yields.

4.2.1 Biphenyl 2-AG Analog Cannabinoid Binding Data

Literature values for the binding affinity of 2-AG to CB1 and CB2 vary over a significant

range. The reported Ki values of 2-AG to CB1 are 100,14 472,15 538,16 and 2400 nM;17 while

values for CB2 are 100,14 1100,18 and 1400 nM.15 Based on these values we were interested in

any compounds that would have binding affinities <1000 nM.

The first two biphenyl analogs synthesized were 18 and 19 (Table 4.1). It was expected

that the fully deprotected 2-MAG, 18, would have the greater binding affinity as it more closely

resembled 2-AG. However, the intermediate product, diglyceride 19, showed binding affinities

towards the CB1 receptor, with and without PMSF, of ~730 nM and ~440 nM respectively,

whereas the glycerol ester had affinities >1000 nM. It was also expected that the FAAH

inhibitor, PMSF, would enhance the binding affinity of any glycerol compounds, as it has been

reported that 2-AG is a substrate for hydrolysis from FAAH.19 With the diglyceride having a

higher affinity towards the CB1 than the glycerol ester, we continued to isolate the intermediate

along with the glycerol product to observe if this trend continued, as the diglycerides may

establish a desired stabilization of the molecule.

Most of the synthesized compounds had CB1 and CB2 binding affinities of >1000 nM.

Besides 18, there were three additional analogs that displayed submicromolar affinities. Fully

saturated biphenyl analog 25 exhibited CB1 Ki values of ~900 and ~780 nM for CB1, with and

without PMSF respectively. Analog 35, with a longer carbon chain between the biphenyl and

ester groups, had the best CB1 binding affinities of ~460 and ~780 nM, and biphenyl ether

analog 42 had CB1 binding affinities of ~700 and >1000 nM. These compounds were

138

reevaluated using 8-pt binding assays, however, complete curves were not obtained, thereby

limiting the reliability of these results.

With 2-AG exhibiting relatively weaker binding affinities to the CB receptors while

being extremely efficacious, solely relying on in vitro binding assays may be misleading. Since

accurate 8-pt binding data was not available, we were curious to observe any in vivo activity of

these ligands before pursuing additional biphenyl type 2-AG analogs. Compounds 18, 19 and 35

were submitted to Dr. Järbe for in vivo studies in mice. Preliminary locomotion analysis studies

were conducted in which mice were dosed with the ligands at 3, 10, and 30 mg/Kg.

Unfortunately, none of these compounds displayed a significant difference in locomotion

compared to vehicle.

Table 4.1 2-point cannabinoid receptor binding assay data Cmpd No. AM Structure rCB1 (nM) CB2 (nM)

untreated w/ PMSF mouse human

17 9076 O

OO

OH

O

~730 ~440 >1000 >1000

18 9077

O

OOH

OH

>1000 >1000 >1000 >1000

19 9080 O

O

OH

O

O

>1000 >1000 >1000 >1000

23 9078 O

O

O

O

OH

>1000 >1000 >1000 >1000

139

24 9075

O

O

OH

OH

~900 ~780 >1000 >1000

30 9087 O

O

O

O

OH

>1000 >1000 >1000 >1000

31 9086 O

O

OH

OH

>1000 >1000 >1000 >1000

34 9089 O

O

O

O

OH

>1000 >1000 >1000 >1000

35 9090 O

O

OH

OH

~460 ~780 >1000 >1000

36 10321

O

O

O

O

OH

>1000 >1000 >1000 >1000

37 10322

O

O

OH

OH

>1000 >1000 >1000 >1000

42 10330

O

O

O

OH

OH

~700 >1000 >1000 >1000

44 10331 O

O

OH

OH

O

>1000 >1000 >1000 >1000

140

4.3 Modification of the Glycerol Head Group

With evidence indicating that biphenyl moieties are not suitable replacements for the

arachidonic tail of 2-AG we shifted our focus to modifications of the glycerol ester in order

reduce the chemical and metabolic instability of this functional group. Other syntheses of 2-AG

analogs indicated that the two hydroxyl groups of the glycerol ester were important for

interactions with the CB receptor. This was indicated by the markedly reduced CB activity when

the hydroxyl groups were replaced with fluorine5 or conformationally constrained in a ring

system.8

We had previously synthesized compounds containing a methyl in the head group to

increase metabolic stability (see Chapter 2), based on its success on other arachidonic analogs.4

Parkkari, et al., reported that α-methylation to the ester showed a slight increase in metabolic

stability, but there was no indication that this would also limit any acyl migration.5 Therefore, we

decided to investigate the addition of methyl groups at the sn-1 and 3 positions to increase

metabolically stablity towards FAAH and MGL, while preventing acyl migration as described in

Chapter 3.

4.3.1 Synthesis of 2,4-dihydroxypentan-3-arachidonoates

Due to the instability of 2-AG, any modifications on the molecule would have to be

completed before the glycerol ester is formed, as reactions performed in the presence of the

glycerol ester may cause acyl migration or hydrolysis. We started with the bromination of acetyl

acetone 45 to give 46, which existed as tautomers, with the keto-enol form favored over the

diketone in the NMR using chloroform-d as the solvent. The keto-enol form is less polar than the

keto form due to intramolecular hydrogen bonding, which is why the keto-enol form is favored

in chloroform. Arachidonic acid was treated with NaH at 0 °C to create sodium arachidonate in

141

situ, which was then treated with 46 and heated to 50 °C. This reaction yielded arachidonic ester

47 in 60% yield (Scheme 4.4). Treatment of diketone 47 with NaBH4 yielded a racemic ±48 as

the major isomer (81%), along with minor isomers 49 (5%) and 50 (14%). The addition of

diethylmethoxyborane to coordinate to the diketones directed a syn-borohydride reduction. This

produced the syn-diols 49 (26%) and 50 (67%) as the major product, with ±48 (7%) as the minor

isomer.

Because the reduction of the diketone using NaBH4 yielded a mix of isomers that still

needed to be separating, other methods were investigated to yield isomers in high enantiomeric

excess. Milder reducing conditions were attempted to yield a β-hydroxy ketone so that the

subsequent reduction of the ketone could be stereochemically controlled. Reactions with

Catecholborane, BH3-THF, and Corey-Bakshi-Shibata catalyst were all attempted, however, no

reduction was observed. These attempted reagents are used to synthesize stereochemically

controlled 1,3-diols from β-hydroxy ketones, and apparently are not suitable for 1,3-diketone

reduction.

Scheme 4.4 Synthesis of to 2,4-dihydroxypentan-3-arachidonoates

O O Br2, H2O, CCl4 O O

Br

OH O

Br

O

OHNaH, DMF0-50 °C

O

OO

O

60%

45 46

46

47

142

O

OOH

OHO

OOH

OHO

OOH

OH

50meso-(2R, 3r, 4S)

49meso-(2R, 3s, 4S)

±48(2R, 4R) & (2S, 4S)

O

OO

O

O

OO

O

NaBH4THF/MeOH-78 °C

Et2BOMeNaBH4THF/MeOH-78 °C

67%26%7%

14%5%81%

~60%

~60%

47

47

4.3.2 Synthesis of 1,3-dihydroxybutan-2-arachidonoates

After synthesizing the dimethyl-glycerols, we were interested in the properties of a

monomethyl-glycerol analog of 2-AG (53, Scheme 4.5). With no methyl-substituted glycerols

available, a keto-oxirane opening was attempted first. Epoxide 52 was synthesized by

epoxidation of α,β-unsaturated ketone 51. Following methodologies for the synthesis of 47,

arachidonic acid was treated with sodium hydride to generate the arachidonic anion in attempt to

open the epoxide ring activated by diethylmethoxyborane to direct the nucleophilic attack.

Unfortunately, no ring opening product was observed. This is most likely due to the conjugate

base of arachidonic acid being a weak nucleophile from the negative charge shared over the two

oxygen molecules.

Scheme 4.5 Attempted keto-oxirane ring opening

O

OH

O

O

OH

ONaH, Et2BOMeTHF, DMFO H2O2, NaHCO3, 3h

O

O

51 52 53

143

Recently, (±)butane-1,2,3-triol (54, Scheme 4.6) became commercially available, which

allowed us to pursue of the chemoenzymatic strategy towards a mono-methyl analog of 2-AG, as

was used in the synthesis of 2-AG. We started with CAL catalyzed 1,3-esterification of 54,

however, the C3 hydroxyl was not esterified by the immobilized lipase. Rather, CAL preferably

esterified the C1 and C2 hydroxyls (56, Scheme 4.6). Alternatively, 3-esterification may have

occurred followed by rapid transesterification. Therefore, chemoenzymatic methods were not

suitable for the generation of 55 based on unsuccessful experiements.

Scheme 4.6 Chemoenzymatic esterification of ±butane-1,2,3-triol

C. antarcticaCH2Cl2OHHO

OH

OOOH

O O

OHOO

O

O

213

213

54

55

56

With the ‘ABA’ triglyceride strategy not readily accessible through this method, the next

attempt was to synthesize an ‘AAA’ (symmetrical) triglyceride and proceed through lipase

catalyzed hydrolysis. An excess of arachidonic acid was coupled to 54 with EDCI and DMAP to

produce ‘AAA’ triglyceride 57 in 96% yield. With the issue of esterifying 54 with C. antarctic,

hydrolysis was expected to be slower than during 2-AG synthesis. Treatment of 57 with CAL

resulted in 30% conversion after 2 h. Due to the slow rate, a second portion of lipase was added

and the reaction continued until a halt in progress was observed. Reaction was complete after 4

h, and 53 was isolated in 73% yield (Scheme 4.7).

144

Scheme 4.7 Successful synthetic strategy towards a monomethyl 2-AG analog

O

OH 54

EDCI, DMAPCH2Cl2

O

O

O

O

O

C19H31

O

C19H31

57

O

O

OH

OH

53

C. antarcticaethanol, 4 h

96%

73%

4.4 Purification and Isolation of 2,4-dihydroxypentan-3-arachidonoate Isomers

With asymmetric synthesis of the 1,3-diols unsuccessful, we then concentrated on the

separation and identification of each individual isomer. From TLC analysis, it appeared that a

chromatographic separation would be possible, but the solvent system would have to be

optimized as it was unclear how many products were formed from each reaction. The NMR

spectrum of the mixture of products indicated that there were three isomers, which was most

apparent from the α-CH2 protons (labeled Z, Figure 4.2). Analysis of protons X and Y indicate

that there were probably two major products and one minor one.

145

Figure 4.2.esp

4.80 4.75 4.70 4.65 4.60Chemical Shift (ppm)

0

0.005

0.010

0.015

0.020

0.025

0.030

0.035

0.040

0.045N

orm

aliz

ed In

tens

ity

Y

Y

O

OOH

OHX

X

YZ

Figure 4.2.esp

4.35 4.30 4.25 4.20 4.15 4.10 4.05 4.00Chemical Shift (ppm)

0

0.005

0.010

0.015

0.020

0.025

0.030

0.035

Nor

mal

ized

Inte

nsity

X

X

O

OOH

OHX

X

YZ

Figure 4.2 1H NMR of 1,3-diol products after NaBH4 reduction of 1,3-diketone

146

Figure 4.2.esp

2.65 2.60 2.55 2.50 2.45 2.40 2.35 2.30Chemical Shift (ppm)

0

0.01

0.02

0.03

0.04

0.05

0.06

0.07

0.08

0.09

Nor

mal

ized

Inte

nsity

Z

Z

Z

O

OOH

OHX

X

YZ

Figure 4.2 1H NMR of 1,3-diol products after NaBH4 reduction of 1,3-diketone

The first attempt at separating the three isomers was performed on an extended column

with a solvent gradient from 0 to 35% ethyl acetate in hexanes. This successfully gave pure

fractions for each isomer, however, most of the material remained impure due to separation

overlaps. This would require additional chromatographic purifications.

To obtain a better separation of the three products, the solvent system needed to be

optimized. From the first solvent system (35% ethyl acetate / hexanes) we moved to a less polar

solvent with hexanes, but at a higher percentage. This led to the trial of using 90% ether /

hexanes and 100% ether. While the products had higher Rf values, the separation was worse

(Figure 4.3). The next solvent attempted was 100% CH2Cl2, which is much more polar than

ether and was expected to increase the Rf of the very polar glycerol products. However,

unexpectedly the products did not move from the baseline. This was thought to occur from an

absence of hydrogen bonding interactions between the glycerol moiety and the CH2Cl2. CH2Cl2

147

was then used in place of hexanes as the base solvent while mixed with 50% ether or 50% ethyl

acetate. This led to better separation of the three products as compared to using hexanes as the

base solvent. The final attempt to optimize separation utilized 10% acetone / CH2Cl2. This

solvent system produced the best separation as the isomers did not overlap, and were most

efficient in obtaining the greatest isolated yields of each pure isomer.

10% acetone in CH2Cl2

50% ethyl acetate in CH2Cl2

50% ether in CH2Cl2

10% hexanes in ether

100% ether 100% CH2Cl2

Figure 4.3 Product separation from TLC analysis of tested solvent systems

4.5 Identification of pent-2,4-ol-3-arachidonoate Stereochemistry

Reduction of 2-4 diketones can result in either syn- or anti-2-4 diols (Figure 4.4). Syn-

diols result in a pseudo-asymmetric carbon at the C3 position. This C3 has a relative chirality

resulting in two meso compounds: an all syn-meso (2R, 3r, 4S) and an 2,3-anti meso (2R, 3s, 4S)

compound. When the 2-4 diols are anti the C3 is no longer pseudo-asymmetric resulting in a

pair of enantiomers (Figure 4.4). Thus the three products observed through TLC and from the

mixed NMR are the racemate ±48, meso-(2R, 3r, 4S) 49, and meso-(2R, 3s, 4S) 50.

148

ROOH

OH

ROOH

OH

ROOH

OH

ROOH

OH

ROOH

OH

ROOH

OH

meso-(2R, 3r, 4S) meso-(2R, 3s, 4S)(2R, 4R) (2S, 4S)

1

32

45

Figure 4.4 Expected products from NaBH4 reduction of a 2-4 diketone

After identification of all possible products as a result of reduction, and the ability to

isolate each product, difficulty lies in distinguishing the different stereoisomers. The first

fraction, with an Rf of 0.42 (10% acetone / CH2Cl2), was the easiest to distinguish because of an

obvious doublet of doublets (dd) at 4.67 ppm (Figure 4.5). The dd splitting indicates the

adjacent protons at the C2 and C4 positions are not equivalent. We know that in (±)48 the C3

proton will have a syn-hydrogen and anti-hydrogen in both enantiomers. The coupling to each of

these should be different, which is why a dd is observed. The mutliplets at 4.23 and 4.08 ppm

also indicate the difference between the relationships of the C2-C3 protons compared to the the

C3-C4 protons.

For the two meso compounds one would expect to observe a triplet for the C3 proton as it

would have an equivalent stereochemical relationship to the C2 and C4 protons. This would

result in a triplet as opposed to the dd observed for the racemic product (±48). As seen in Figure

4.5, both 49 and 50 display clean triplets at 4.68 and 4.73 ppm respectively.

149

+-48.esp

4.65 4.60 4.55 4.50 4.45 4.40 4.35 4.30 4.25 4.20 4.15 4.10 4.05Chemical Shift (ppm)

0

0.05

0.10

Nor

mal

ized

Inte

nsity

0.760.750.73

M02(dd) M03(quind) M04(m)

4.67

4.67

4.66

4.66

4.25

4.25

4.24

4.24

4.23

4.22

4.22

4.21

4.20

4.20

4.11

4.10

4.09

4.07

4.06

4.05

O

OOH

OH

±48

49.esp

4.70 4.65 4.60 4.55 4.50 4.45 4.40 4.35 4.30 4.25 4.20 4.15 4.10 4.05 4.00 3.95Chemical Shift (ppm)

0

0.05

0.10

0.15

Nor

mal

ized

Inte

nsity

1.950.92

J(M01)=6.41 Hz

J(M01)=6.41 Hz

M01(t)

M03(m)

4.02

4.03

4.03

4.04

4.67

4.68

4.70

O

OOH

OH

49

Figure 4.5 Expansion of splitting patterns of C2, C3, and C4 protons

150

50.esp

4.80 4.75 4.70 4.65 4.60 4.55 4.50 4.45 4.40 4.35 4.30 4.25 4.20 4.15 4.10 4.05Chemical Shift (ppm)

0

0.025

0.050

0.075

Nor

mal

ized

Inte

nsity

2.000.95

J(M01)=3.05 Hz

J(M01)=3.05 Hz

M01(t)

M02(td)

4.11

4.12

4.12

4.13

4.14

4.14

4.72

4.73

4.74

O

OOH

OH

50

Figure 4.5 Cont’d Expansion of splitting patterns of C2, C3, and C4 protons

To correctly assign 49 and 50 to the proper meso compounds, one must closely examine

the coupling constants. One triplet has a J2-3 and J3-4 of 6.41 Hz, and the other 3.05 Hz. If we

visualize the Newman projections of 49 and 50, the simplest forms, based on their

stereochemistry, show the dihedral angle of HA-HB to be 180° for the meso-(2R, 3s, 4S) and 60°

for the meso-(2R, 3r, 4S). Based on studies of Karplus20 and Stiles, et al.,21 one would associate

a dihedral angle of 60° with a smaller coupling constant than one from a dihedral angle of 180°

(Figure 4.6). Based on these observations it is speculated that the second eluting product (Rf =

0.35, 10% acetone / CH2Cl2), with a JAB of 6.10 Hz, is associated with meso-(2R, 3s, 4S) 49.

The last eluting product (Rf = 0.25, 10% acetone / CH2Cl2), with a JAB of 3.05 Hz, is speculated

to be meso-(2R, 3r, 4S) 50.

151

HBR

HHOHAHO

RHB

HHOHAHO

22 334 4

OHHA

C4

HB R

OHHA

C4

R HB

O

OOH

OH2

4

3O

OOH

OH2

4

3

60°

180°3.05 Hz6.41 Hz

49 50

C2-C3 C2-C3

meso-(2R, 3s, 4S) meso-(2R, 3r, 4S)

Figure 4.6 Newman projects of the two meso compounds: 49 & 50

4.6 Chemically and Metabolically Stable 2-AG Analog Data

4.6.1 Chemical Stability of 2-AG Analogs

As described previously, the tendency of 2-AG to undergo facile acyl migration can be a

hindrance when preparing large quantities, or storing over long periods of time. However, it has

been observed (through NMR) that the initial ratio of 2-AG to 1-AG has a large effect on the rate

at which acyl migration occurs. Very pure samples are better suited for long term storage, while

impure samples initiate acyl migration at higher rates. A sample of 2-AG (>98%), sent for use in

pharmacological studies, was stored in ethanol at -80 °C while being warmed to room

temperature and re-cooled numerous times. After six weeks this same sample was analyzed by

NMR and the sample was 92% 2-AG compared to the 98% after synthesis. However, this

observation only indicates that purer samples of 2-AG are fairly stable in storage, and provided

152

no indication of the tendency for acyl migration when 2-AG is subjected to in vitro assays or

when administered in vivo.

While the ratio of 2-AG to 1-AG can vary slightly for each synthesis, crude NMR of

NaBH4 reductions of 47 to ±48, 49, and 50 indicated no formation of a 1-AG variant after

reaction (2 h). However, 1-AG analogs (±48, 49, and 50) were isolated when the reaction was

allowed to occur overnight and also when the reaction was neutralized with citric acid as

opposed to hydrochloric acid. Even if a minimal amount of the 1-AG variant of these analogs

was observed, column chromatography was able to separate them, as the 1-AG dimethyl analog

could be separated from the desired compound; whereas, natural 1-AG and 2-AG are not

separable by column chromatography.

4.6.2 Metabolic Stability of 2-AG Analogs Compared to 2-AG

To compare the metabolic stability of our synthesized 2-AG analogs to that of 2-AG, we

performed a hMGL substrate assay. This assay exposed 2-AG, (±)48, 49, 50, and 53 to hMGL

and measured the concentration of the initial ligand and arachidonic acid produced from

metabolic hydrolysis after 30 minutes (Table 4.2).

The important data was the accumulation of AA after 30 minutes of incubation with

hMGL. 2-AG was almost fully metabolized by the 30 min with 86% AA accumulation. The

graph shows that the majority of 2-AG is hydrolyzed by hMGL within 3-5 minutes. Analog

(±)48 appeared to be the most metabolically stable compound as there was no measurable AA

measured after 30 m. The 2,3-anti analog 49 exhibited 18% AA accumulation, and the 2,3-cis

analog 50 was almost 3x more stable with only 6.6% AA accumulated. Mono-methyl analog 53

was, as expected, not as stable as the dimethyl analogs, but still more metabolically stable than 2-

AG. An interesting observation after 30 m was the reduction in the amount of original substrate

153

remaining while a non-proportional amount of AA accumulated. It is postulated that the

formation of an enzyme-substrate complex occurred where the ligand entered the binding

domain of hMGL but was not hydrolyzed, evident by the lack of AA formation, and may behave

as an inhibitor.

Table 4.2 hMGL substrate assay results

Compound AM After 30 minutes

% Substrate remaining

% AA accumulated

2-AG 2-AG 6.0 86 ±48 10336 52 0.0 49 10335 51 18 50 10334 65 6.6 53 10342 15 22

0

50

100

150

200

0 5 10 15 20 25 30 35

Conc

entr

atio

n (u

M)

Time (minutes)

2-AG Stability with hMGL

[2-AG] (uM)

[AA] (uM)

-50

0

50

100

150

200

250

0 5 10 15 20 25 30 35

Conc

entr

atio

n (u

M)

Time (minutes)

±48 (AM-10336) Stability with hMGL

[AM-10336] (uM)

[AA] (uM)

154

4.6.3 Cannabinoid In Vitro Binding Assay Data

For in vitro binding data we were able to measure the inhibition of cAMP release for

(±)48, 49, and 50. The data are summarized in Table 4.3. Compounds (±)48, 49, and 50 all

-50

0

50

100

150

200

250

0 5 10 15 20 25 30 35

Conc

entr

atio

n (u

M)

Time (minutes)

49 (AM-10335) Stability with hMGL

[AM-10335] (uM)

[AA] (uM)

-50

0

50

100

150

200

250

0 5 10 15 20 25 30 35

Conc

entr

atio

n (u

M)

Time (minutes)


[AM-10334] (uM)

[AA] (uM)

0

50

100

150

200

250

0 5 10 15 20 25 30 35

Conc

entr

atio

n (u

M)

Time (minutes)


[AM-10342] (uM)

[AA] (uM)

155

behaved as agonists in the cAMP assay with 67%, 49%, and 58% decrease in stimulation,

respectively.

Table 4.3 Cyclic AMP assay for 2-AG dimethyl analogs

Compound AM rCB1 IC50 (μM) 2-AG 0.010 (±)48 10336 1.3

49 10335 1.3 50 10334 2.6

4.6.4 Pharmacological Data of 2-AG, 1-AG, AA, and 2-AG Analogs In Vivo

Compounds (±)48 and 50 were submitted for a pharmacological comparison to 2-AG.

Locomotor and rearing activity were examined in mice, dosed with 2-AG, 1-AG, arachidonic

acid, (±)48, and 50. The locomotion test uses open-field observation of movement in mice after

being dosed, while rearing is the tendency for the mice to stand on its hind legs after

administration.

The dose-dependent response curves for locomotion (Figure 4.7) and rearing (Figure 4.8)

of 2-AG compared to (±)48 and 50, exhibit significantly different profiles. Dosing of 2-AG

showed an immediate attenuation of locomotion (and rearing) in mice after 15 minutes,however,

this effect dissipated after 30 minutes. When observing (±)48 and 50 in the same experiment, the

subjects experienced a delayed onset of action from these analogs that was not observed with 2-

AG. The 30 mg/kg dose of (±)48 had a similar effect at 30 minutes as the 10 mg/kg dose of 2-

AG produced at 15 minutes (first reading).

To establish whether these observed effects are CB1 mediated, compounds were

administered along with the CB1 inverse agonist SR141716 (SR, Rimonabant). If an attenuation

of action was observed then it would indicate the response was an effect of CB1 activation.

Administering 2-AG with SR showed no change in the pharmacological profile (data not shown).

156

On the other hand, treatment of 10 mg/kg of SR with 30 mg/kg of (±)48 attenuated the response

to that of the 10 mg/kg dose of (±)48. This implied that the effects of (±)48 were a result of

ligand interactions with the CB1 receptor.

Figure 4.7 Locomotor activity following administration of 2-AG analogs (All data is obtained from one single experiment).

0

200

400

600

800

1000

1200

1400

1600

1800

2000

15 30 45 60

Measured Bin Time (min)

Locomotion in 2-AG Dosed Mice

Vehicle

1mg/kg 2-AG

3mg/kg 2-AG

10mg/kg 2-AG

0.00

200.00

400.00

600.00

800.00

1000.00

1200.00

1400.00

1600.00

1800.00

2000.00

15 30 45 60


Locomotion with 2-AG Analogs

Vehicle

10mg/kg ±48

30mg/kg ±48

30mg/kg ±48+10mg/kgSR

10mg/kg 50

157

Figure 4.8 Rearing activity following administration of 2-AG analogs (All data is obtained from one single experiment).

The CB1 inverse agonist, SR, did not attenuate the pharmacological effects of 2-AG.

This implied that the observed pharmacological profile may not be strictly from 2-AG, if at all.

We know that 2-AG did not go through acyl migration prior to injection; yet it is unknown

0

20

40

60

80

100

120

15 30 45 60


Rearing in 2-AG Dosed Mice

Vehicle

1mg/kg 2-AG

3mg/kg 2-AG

10mg/kg 2-AG

0

20

40

60

80

100

120

15 30 45 60


Rearing with 2-AG Analogs

Vehicle

10mg/kg ±48

30mg/kg ±48

30mg/kg ±48+10mg/kgSR

10mg/kg 50

158

whether this migration can occur in vivo. To resolve this issue, mice were given a dosage of 1-

AG (3 mg/kg) and the results were compared to those for 2-AG. Figure 4.9 illustrates that the

profiles of 1-AG and 2-AG at 3 mg/kg very similar in regards to locomotion and rearing. With

1-AG and 2-AG exhibiting similar pharmacological profiles, this suggests that a metabolite of

these compounds may be causing these effects and acting somewhere other than the CB

receptors as SR is unable to attenuate any of these effects.

It was thought that 1-AG and 2-AG were immediately hydrolyzed by MGL post

administration. If this were the case, glycerol and arachidonic acid would be present as

byproducts of hydrolysis. Therefore, mice were administered arachidonic acid and the profile

was compared to 1-AG and 2-AG. The effects of arachidonic acid resembled those of 1-AG and

2-AG, leaving one unable to draw any conclusions regarding the biological effects observed.

The next experiments will have to include MGL inhibitors as well as the administration of 2-AG

to possible understand any pharmacological data pertaining to effects caused by 2-AG.

0

200

400

600

800

1000

1200

1400

1600

1800

2000

15 30 45 60


Locomotion in 2-AG Dosed Mice

Vehicle

1mg/kg 2-AG

3mg/kg 2-AG

10mg/kg 2-AG

3mg/kg 1-AG

10mg_kg_ArachidonicA

3mg_kgArachidonicA

159

Figure 4.9 Locomotor and rearing activity following administration of 2-AG, 1-AG, and arachidonic acid (All data is obtained from a single experiment).

4.7 Conclusion

The study of the true pharmacological properties of 2-AG can be difficult. Administration

of 1-AG, 2-AG, and AA showed similar profiles when administered in vivo, suggesting that the

observed results were not an accurate representation of the pharacodynamics of 2-AG. It is

believed this occurs due to the rapid metabolism from MGL. Therefore, we developed a strategy

to synthesize chemically and metabolically stable 2-AG analogs that would exhibit a longer

duration of action in order to observe a true pharmacological profile.

Our first attempt at synthesizing ligands where the hydrophobic unsaturated arachidonic

tail was replaced with various biphenyl moieties, improved chemical stability but did not render

the ligand resistant to hydrolysis. Through in vitro and in vivo experiments it was determined

these compounds did not display activity and efficacy similar to that of 2-AG. A second strategy

was to modify the glycerol head group and leave the unsaturated arachidonic tail intact. This

was successful in providing chemical and metabolic stability, where no acyl migration was

observed and hydrolysis was drastically reduced (even eliminated in the case of (±)48).

0

20

40

60

80

100

120

140

15 30 45 60


Rearing in 2-AG Dosed Mice Vehicle

1mg/kg 2-AG

3mg/kg 2-AG

10mg/kg 2-AG

3mg/kg 1-AG

10mg_kg_ArachidonicA

3mg_kgArachidonicA

160

In pharmacological studies, it was observed that the 2-AG analogs exhibited a different

profile when observing locomotion. The stable analogs did not have an immediate onset of

action and the effects appeared to last longer than that observed with 2-AG. The effects of the

analogs were also noted to be CB mediated, as an antagonist was able to attenuate the observed

effects. This makes ±48 a promising candidate as a 2-AG replacement and tool to understand the

endogenous effects of the lipid signaling compound.

In summary, the addition of chiral groups into the glycerol ester head-group of 2-AG is a

viable strategy to overcome the chemical and metabolic instabilities observed in 2-AG. It also

appears that the structural moieties that stray from the arachidonic acid tail are not well tolerated

in the CB receptors. While (±)48 appeared to be a chemically and metabolically stable analog of

2-AG, its effects compared the endogenous analog are not fully understood as pharmacological

profiles of 2-AG are not fully understood.

161

4.8 Experimental

O

O

(E)-methyl 3-(biphenyl-4-yl)acrylate, 12. Methyl(triphenylphosphoranylidene)acetate (2.81 g,

8.39 mmol) was added to a solution of biphenyl-4-carbaldehyde (1.52 g, 8.39 mmol) in 20 mL of

anhydrous CH2Cl2 at 0 °C. The reaction was allowed to warm to room temperature and stir

overnight. The solvent was evaporated off under reduced pressure and the residue was

chromatographed on silica gel to yield 12 (1.012 g, 51%) as a white solid. Rf = 0.52 (20% ethyl

acetate / hexanes).

O

OH

(E)-3-(biphenyl-4-yl)acrylic acid, 13. A 1.0M solution of LiOH (0.88 mL, 0.88 mmol) was

added to a solution of (Z)-methyl 3-(biphenyl-4-yl)acrylate (105 mg, 0.44 mmol) in 4mL of

THF. The reaction was allowed to stir under argon for 48 hours. Upon completion, a 1.0M

solution of HCl was added until the reaction mixture was slightly acidic. The product was

extracted with Et2O washed with water, brine, and dried over MgSO4. The organic layer was

concentrated to yield 13 (78 mg, 78%) as a white solid. 1H NMR (500 MHz, CHLOROFORM-

d) δ ppm 7.73 (d, J=7.81 Hz, 2 H) 7.58 - 7.64 (m, 4 H) 7.45 (t, J=7.57 Hz, 2 H) 7.37 (s, 1 H)

7.10 (d, J=12.70 Hz, 1 H) 6.01 (d, J=12.70 Hz, 1 H).

162

O

OH

(E)-4-(biphenyl-4-yl)but-3-enoic acid, 14. A 1.0M solution of potassium tert-butoxide in THF

(1.1 mL, 1.1 mmol) was added to a suspension of (2-carboxyethyl) triphenylphosphonium

bromide (342 mg, 0.83 mmol) in 5 mL of anhydrous THF at -78°C. After 10 minutes biphenyl-4-

carbaldehyde (100 mg, 0.55 mmol) was added portion wise. The reaction was allowed to warm

slowly to room temperature. The reaction was quenched with 1.0M HCl (1.0 mL) and diluted

with ether. The organic layer was separated, dried over MgSO4, and concentrated. The residue

was chromatographed on silica gel to yield 14 (60 mg, 46%) as a white solid. Rf = 0.25 (70%

ethyl acetate / hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.54 - 7.62 (m, 4 H)

7.41 - 7.48 (m, 4 H) 7.31 - 7.37 (m, 1 H) 6.57 (d, J=16.11 Hz, 1 H) 6.30 - 6.39 (m, 1 H) 3.34 (d,

J=6.84 Hz, 2 H).

O

OO

O

O

O

(E)-2-(3-(biphenyl-4-yl)acryloyloxy)propane-1,3-diyl dibutyrate, 15. EDCI (220 mg, 1.15

mmol), DMAP (6 mg, 0.046 mmol) and 1,3-dibutyrate glycerol (106 mg, 0.46mmol) were added

to a solution of 13 (52 mg, 0.23 mmol) in 5mL of anhydrous THF/DCM (1:1). The reaction was

allowed to stir under argon for 4 hours. Upon completion the reaction was diluted with CH2Cl2

and water. The organic layer was concentrated and the residue was chromatographed on silica

gel to yield 15 (85 mg, 84%) as a white solid. Rf = 0.22 (20% ethyl acetate/hexane). 1H NMR

(500 MHz, CHLOROFORM-d) δ ppm 7.76 (d, J=15.63 Hz, 1 H) 7.54 - 7.68 (m, 6 H) 7.46 (t,

163

J=7.57 Hz, 2 H) 7.36 - 7.40 (m, 1 H) 6.49 (d, J=16.11 Hz, 1 H) 4.25 - 4.38 (m, 4 H) 2.30 (t, J =

7.32 Hz, 4H), 1.65 (sxt, J = 7.30 Hz, 4H), 0.95 (t, J = 7.57 Hz, 6H).

O

O

O

O

O

O

(E)-2-(4-(biphenyl-4-yl)but-3-enoyloxy)propane-1,3-diyl dibutyrate, 16. The procedure for

15 was followed to synthesize 16 (93 mg, 83%) as an oil. Rf = 0.64 (35% ethyl acetate /

hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ 7.54 - 7.62 (m, 4H), 7.41 - 7.47 (m, 4H),

7.34 (s, 1H), 6.54 (d, J = 15.63 Hz, 1H), 6.32 (td, J = 7.3, 15.6 Hz, 1H), 5.25 - 5.35 (m, 1H), 4.32

(ddd, J = 3.9, 7.8, 18.0 Hz, 2H), 4.18 (ddd, J = 5.8, 12.2, 19.0 Hz, 4H), 3.29 (dd, J = 1.2, 7.1 Hz,

2H), 2.26 - 2.34 (m, 4H), 1.60 - 1.70 (m, 4H), 0.90 - 0.98 (m, 6H)

O

OO

OH

O

(E)-2-(3-(biphenyl-4-yl)acryloyloxy)-3-hydroxypropyl butyrate, 17, AM9076. Candida

antarctica (150 mg) was added to a solution of 15 (150 mg, 0.39 mml) stirred at room

temperature in anhydrous ethanol (1 mL). The reaction was stirred for 75 minutes. After

consumption of starting material the reaction was halted and was filtered and diluted with

CH2Cl2. The organic layer was removed under reduced pressure and the residue was

chromatographed silica gel to yield 17 (10 mg, 18%) as a white solid. Rf = 0.48 (40% ethyl

acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ 7.76 (d, J = 16.11 Hz, 1H), 7.54 -

7.69 (m, 6H), 7.40 - 7.48 (m, 2H), 7.34 - 7.40 (m, 1H), 6.50 (d, J = 16.11 Hz, 1H), 5.24 (quin, J

= 5.01 Hz, 1H), 4.36 - 4.41 (m, 2H), 3.81 - 3.85 (m, 2H), 2.34 (t, J = 7.32 Hz, 2H), 1.61 - 1.70

164

(m, 2H), 0.93 - 0.98 (m, 3H). 13C NMR (126 MHz, CHLOROFORM-d) δ 174.0, 166.8, 145.7,

140.3, 129.2 (2C), 129.0 (5C), 128.2, 127.8 (2C), 127.3 (2C), 72.8, 62.4, 61.9, 36.2 (3C).

O

OOH

OH

(E)-1,3-dihydroxypropan-2-yl 3-(biphenyl-4-yl)acrylate, 18, AM9077. The reaction

producing 17 also produced 18 (34 mg, 49%) as a white solid. Rf = 0.36 (40% ethyl acetate /

hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ 7.78 (d, J = 16.1 Hz, 1H), 7.54 - 7.67 (m,

6H), 7.42 - 7.51 (m, 2H), 7.34 - 7.41 (m, 1H), 6.54 (d, J = 15.6 Hz, 1H), 5.08 (quin, J = 4.7 Hz,

1H), 3.94 (t, J = 5.1 Hz, 4H), 2.26 (t, J = 5.9 Hz, 2H). 13C NMR (126 MHz, CHLOROFORM-d)

δ 167.4, 145.8, 143.6, 129.2 (2C), 129.0 (5C), 128.2, 127.8 (2C), 127.3 (2C), 80.2, 62.9 (2C).

O

O

O

O

OH

(E)-1-(butyryloxy)-3-hydroxypropan-2-yl 4-(biphenyl-4-yl)but-3-enoate, 19, AM9080. The

procedure for 17 was used synthesize 19 (23 mg, 35%) as a colorless oil. Rf = 0.65 (50% ethyl

acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ 7.52 - 7.62 (m, 4H), 7.40 - 7.47

(m, 4H), 7.31 - 7.37 (m, 1H), 6.55 (d, J = 15.6 Hz, 1H), 6.33 (td, J = 6.4, 16.1 Hz, 1H), 5.14

(quin, J = 5.0 Hz, 1H), 4.34 - 4.39 (m, 1H), 4.22 - 4.31 (m, 1H), 3.75 - 3.78 (m, 2H), 3.32 (dd, J

= 1.2, 7.1 Hz, 2H), 2.30 (t, J = 7.6 Hz, 2H), 2.02 - 2.20 (m, 1H), 1.60 - 1.69 (m, 2H), 0.92 (t, J =

7.6 Hz, 3H). 13C NMR (100 MHz, CHLOROFORM-d) δ 173.8, 171.4, 140.7, 135.9, 129.0 (2C),

127.6 (5C), 127.5, 127.2 (2C), 127.0 (2C), 72.9, 62.9, 62.1, 61.7, 38.6, 18.6, 13.9.

165

O

O

methyl 3-(biphenyl-4-yl)propanoate, 20. 10% Pd/C (50 mg) was added to a solution of 12 (800

mg, 3.35 mmol) in 20 mL of ethyl acetate and was evacuated and purged with H2 (3x). Upon

completion the reaction mixture was filtered and the solvent was removed under reduced

pressure to yield methyl 20 (663 mg, 82%) as a white crystalline. Rf = 0.73 (40% ethyl acetate /

hexane). 1H NMR (500 MHz, CHLOROFORM-d) d ppm 7.50 - 7.60 (m, 4 H) 7.43 (t, J=7.81

Hz, 2 H) 7.31 - 7.36 (m, 1 H) 7.28 (d, J=7.81 Hz, 2 H) 3.69 (s, 3 H) 3.00 (t, J=7.81 Hz, 2 H) 2.68

(t, J=7.81 Hz, 2 H).

O

OH

3-(biphenyl-4-yl)propanoic acid, 21. The procedure for 13 was followed to synthesize 21 (353

mg, 63%) as a white solid. Rf = 0.30 (50% ethyl acetate / hexane). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 7.55 (dd, J=21.97, 7.81 Hz, 4 H) 7.43 (t, J=7.57 Hz, 2 H) 7.33 (t,

J=7.81 Hz, 1 H) 7.29 (d, J=8.30 Hz, 2 H) 3.01 (t, J=7.81 Hz, 2 H) 2.74 (t, J=7.81 Hz, 2 H).

O

O

O

O

O

O

2-(3-(biphenyl-4-yl)propanoyloxy)propane-1,3-diyl dibutyrate, 22. The procedure for 15 was

followed to synthesize 22 (416 mg, 61%) as an oil. Rf = 0.67 (35% ethyl acetate / hexane). 1H

166

NMR (500 MHz, CHLOROFORM-d) δ ppm 7.50 - 7.59 (m, 4 H) 7.43 (t, J=7.57 Hz, 2 H) 7.30 -

7.36 (m, 1 H) 7.26 - 7.29 (m, 2 H) 5.23 - 5.33 (m, 1 H) 4.25 - 4.34 (m, 2 H) 4.09 - 4.20 (m, 2 H)

2.99 (td, J=7.81, 3.42 Hz, 2 H) 2.66 - 2.73 (m, 2 H) 2.25 - 2.31 (m, 4 H) 1.58 - 1.68 (m, 4 H)

0.93 (td, J=7.45, 4.64 Hz, 6 H).

O

O

O

O

OH

2-(3-(biphenyl-4-yl)propanoyloxy)-3-hydroxypropyl butyrate, 23, AM9078. The procedure

for 17 was followed to synthesize 23 (165 mg, 59%) as a white solid. Rf = 0.46 (40% acetone /

hexane). 1H NMR (500 MHz, CHLOROFORM-d) δ 7.49 - 7.59 (m, 4H), 7.41 (t, J = 7.6 Hz,

2H), 7.29 - 7.34 (m, 1H), 7.22 - 7.28 (m, 2H), 5.08 (quin, J = 5.13 Hz, 1H), 4.34 (dd, J = 4.4,

11.7 Hz, 1H), 4.23 (dd, J = 5.9, 11.7 Hz, 1H), 3.63 - 3.70 (m, 2H), 2.98 (t, J = 7.6 Hz, 2H), 2.65 -

2.71 (m, 2H), 2.46 (t, J = 5.6 Hz, 1H), 2.30 (t, J = 7.6 Hz, 2H), 1.60 - 1.68 (m, 2H), 0.93 (t, J =

7.1 Hz, 3H). 13C NMR (100 MHz, CHLOROFORM-d) δ 173.6, 173.1, 141.1, 139.6, 129.0,

129.0, 127.5, 127.4, 127.2, 72.2, 62.7, 61.5, 36.4, 35.8, 30.7, 18.7, 13.8.

O

O

OH

OH

1,3-dihydroxypropan-2-yl 3-(biphenyl-4-yl)propanoate, 24, AM9075. The reaction producing

23 also resulted in 24 (100 mg, 35%) as a colorless oil. Rf = 0.14 (40% acetone / hexane). 1H

NMR (399 MHz, CHLOROFORM-d) δ 7.53 - 7.58 (m, 2H), 7.47 - 7.53 (m, 2H), 7.37 - 7.45 (m,

2H), 7.29 - 7.35 (m, 1H), 7.23 - 7.28 (m, 2H), 4.89 (quin, J = 4.95 Hz, 1H), 3.73 (d, J = 4.40 Hz,

167

4H), 2.99 (t, J = 7.33 Hz, 2H), 2.77 (br. s., 1H), 2.72 (t, J = 8.06 Hz, 2H). 13C NMR (100 MHz,

CHLOROFORM-d) δ 173.4, 141.0, 139.6, 139.5, 129.0, 129.0, 127.5, 127.5, 127.2, 75.3, 62.2,

36.0, 30.8.

O

O

methyl 6-(biphenyl-4-yl)hex-5-ynoate, 27. Pd(PPh3)2Cl2 (147 mg, 0.21 mmol), copper iodide

(40 mg, 0.21 mmol) and triethylamine (1.2 mL, 8.6 mmol) were added to a solution of 4-

bromobiphenyl (1.0 g, 4.3 mmol) and methyl 5-hexynoate (561 μL, 4.3 mmol) in 7 mL of

anhydrous DMF under an atmosphere of argon at 65 °C. The reaction was allowed to stir for

18hrs, diluted with Et2O and quenched with water. The organic layer was separated, dried over

NaSO4 and concentrated. The resulting residue was chromatographed on silica gel to yield 27

(742 mg, 62%) as an oil. Rf = 0.70 (35% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 7.58 (dd, J=7.3, 1.0 Hz, 2 H) 7.50 - 7.54 (m, 2 H) 7.41 - 7.48 (m, 4

H) 7.31 - 7.38 (m, 1 H) 3.66 - 3.71 (m, 3 H) 2.46 - 2.58 (m, 4 H) 1.95 (quin, J=7.2 Hz, 2 H) -0.04

- 0.02 (m, 1 H).

O

OH

6-(biphenyl-4-yl)hex-5-ynoic acid, 28. The procedure for 13 was followed to synthesize 28 (641

mg, 91%) as a white solid. Rf = 0.25 (25% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 7.55 - 7.60 (m, 2 H) 7.49 - 7.54 (m, 2 H) 7.40 - 7.48 (m, 4 H) 7.31 -

7.38 (m, 1 H) 2.58 (t, J=7.3 Hz, 2 H) 2.54 (t, J=7.1 Hz, 2 H) 1.96 (quin, J=7.1 Hz, 2 H).

168

O

O

O

O

O

O

2-(6-(biphenyl-4-yl)hex-5-ynoyloxy)propane-1,3-diyl dibutyrate, 29. The procedure for 15

was followed to synthesize 29 (930 mg, 80%) as a colorless oil. Rf = 0.67 (25% ethyl acetate /

hexanes). 1H NMR (399 MHz, CHLOROFORM-d) δ ppm 7.58 (d, J=7.3 Hz, 2 H) 7.50 - 7.55

(m, 2 H) 7.40 - 7.49 (m, 4 H) 7.32 - 7.38 (m, 1 H) 5.25 - 5.35 (m, 1 H) 4.33 (dd, J=12.1, 4.0 Hz,

2 H) 4.17 (dd, J=11.7, 5.9 Hz, 2 H) 2.53 (dt, J=12.5, 7.0 Hz, 4 H) 2.31 (t, J=7.7 Hz, 4 H) 1.95

(quin, J=7.3 Hz, 2 H) 1.65 (sxt, J=7.3 Hz, 5 H) 0.94 (t, J=8.1 Hz, 6 H).

O

O

O

O

OH

1-(butyryloxy)-3-hydroxypropan-2-yl 6-(biphenyl-4-yl)hex-5-ynoate, 30, AM9087. The

procedure for 17 was followed to synthesize 30 (27 mg, 79%) as a colorless oil. Rf = 0.30 (35%


7.50 - 7.55 (m, 2 H) 7.40 - 7.48 (m, 4 H) 7.32 - 7.38 (m, 1 H) 5.12 (quin, J=5.0 Hz, 1 H) 4.35

(dd, J=12.0, 4.6 Hz, 1 H) 4.25 (dd, J=12.2, 5.9 Hz, 1 H) 3.70 - 3.78 (m, 2 H) 2.58 (t, J=7.3 Hz, 2

H) 2.53 (t, J=6.8 Hz, 2 H) 2.32 (t, J=7.3 Hz, 2 H) 2.09 (br. s., 1 H) 1.96 (quin, J=7.2 Hz, 2 H)

1.63 - 1.70 (m, 2 H) 0.94 (t, J=7.6 Hz, 3 H). 13C NMR (100 MHz, CHLOROFORM-d) δ 173.6,

173.1, 132.2, 129.1, 127.7, 127.2, 127.1, 122.8, 89.5, 81.7, 71.6, 62.2, 61.7, 36.2, 33.3, 24.1,

19.1, 18.6, 13.9.

169

O

O

OH

OH

1,3-dihydroxypropan-2-yl 6-(biphenyl-4-yl)hex-5-ynoate, 31, AM9086. Candida antarctica

(100 mg) was added to a solution of 29 (80 mg, 0.17 mmol) stirred in anhydrous ethanol (1 mL)

under an atmosphere of argon. After starting material was consumed additional lipase (100 mg)

was added. After an additional 1h the reaction was diluted with CH2Cl2 and the lipase was

filtered off. The solvent was removed under reduced pressure, and the resulting residue was

chromatographed on methanol washed silica gel to yield 31 (44 mg, 78%) as white solid. Rf =

0.28 (70% ethyl acetate / hexanes). mp 82-84 °C. 1H NMR (399 MHz, CHLOROFORM-d) δ

ppm 7.58 (d, J=8.1 Hz, 2 H) 7.50 - 7.55 (m, 2 H) 7.40 - 7.49 (m, 4 H) 7.31 - 7.38 (m, 1 H) 4.95

(dt, J=9.3, 4.5 Hz, 1 H) 3.84 (br. s., 4 H) 2.61 (t, J=7.7 Hz, 2 H) 2.53 (t, J=7.0 Hz, 2 H) 2.22 (br.

s., 2 H) 1.97 (dt, J=14.1, 7.2 Hz, 2 H). 13C NMR (100 MHz, CHLOROFORM-d) δ 173.6, 140.7,

140.6, 132.2, 129.1, 127.8, 127.2, 89.6, 81.7, 62.7, 33.4, 24.1, 19.1.

O

O

O

O

O

O

2-(6-(biphenyl-4-yl)hexanoyloxy)propane-1,3-diyl dibutyrate, 32. The procedure for 20 was

followed to synthesize 32 (100 mg, 76%) as a colorless oil. Rf = 0.59 (35% ethyl acetate /


H) 7.39 - 7.46 (m, 2 H) 7.29 - 7.35 (m, 1 H) 7.24 (d, J=8.1 Hz, 2 H) 5.23 - 5.31 (m, 1 H) 4.30

170

(dd, J=11.7, 4.4 Hz, 2 H) 4.15 (dd, J=11.7, 5.9 Hz, 2 H) 2.65 (t, J=8.1 Hz, 2 H) 2.26 - 2.37 (m, 6

H) 1.59 - 1.74 (m, 8 H) 1.35 - 1.48 (m, 2 H) 0.95 (t, J=7.3 Hz, 6 H).

O

O

O

O

O

O

(Z)-2-(6-(biphenyl-4-yl)hex-5-enoyloxy)propane-1,3-diyl dibutyrate, 33. To a solution of

Ni(OAc)2•4H2O (373 mg, 1.5 mmol) in anhydrous methanol (10 mL), was added NaBH4 (64

mg, 1.7 mmol) at room temperature under and an argon atmosphere. This mixture was

immediately put under vacuum and purged with H2 (3 times) and allowed to stir for 5 minutes.

The solution was treated with ethylenediamine (147 μL, 2.2 mmol) and stirred for an additional 5

minutes, at which, 29 (400 mg, 0.85 mmol) in 5mL of anhydrous methanol was added. The

mixture was stirred at room temp under an atmosphere of H2 for 2 h. The reaction mixture was

filtered through celite, and the filtrate was diluted with Et2O and brine. The organic phase was

separated and the aqueous phase was extracted 5 times with Et2O, and the combined organic

layers were washed dried over MgSO4. The resulting solution was concentrated and

chromatographed on silica gel to yield 33 (319 mg, 79%) as a colorless oil. Rf = 0.54 (25% ethyl

acetate / hexanes). 1H NMR (399 MHz, CHLOROFORM-d) δ ppm 7.59 (dd, J=12.5, 8.1 Hz, 4

H) 7.44 (t, J=7.7 Hz, 2 H) 7.31 - 7.38 (m, 3 H) 6.49 (d, J=11.7 Hz, 1 H) 5.66 (dt, J=11.5, 7.1 Hz,

1 H) 5.22 - 5.31 (m, 1 H) 4.29 (dd, J=12.1, 4.0 Hz, 2 H) 4.14 (dd, J=11.7, 5.9 Hz, 2 H) 2.44 (q,

J=7.3 Hz, 2 H) 2.38 (t, J=7.7 Hz, 2 H) 2.28 (t, J=7.3 Hz, 4 H) 1.82 (quin, J=7.5 Hz, 2 H) 1.63

(sxt, J=7.5 Hz, 4 H) 0.93 (t, J=7.7 Hz, 6 H).

171

O

O

O

O

OH

1-(butyryloxy)-3-hydroxypropan-2-yl 6-(biphenyl-4-yl)hexanoate, 34, AM9089. The



7.50 - 7.53 (m, 2 H) 7.40 - 7.45 (m, 2 H) 7.30 - 7.34 (m, 1 H) 7.24 (d, J=8.3 Hz, 2 H) 5.09 (quin,

J=5.1 Hz, 1 H) 4.32 (dd, J=11.2, 4.9 Hz, 1 H) 4.23 (dd, J=11.7, 5.9 Hz, 1 H) 3.72 (dd, J=4.6, 1.7

Hz, 2 H) 2.66 (t, J=7.8 Hz, 2 H) 2.37 (t, J=7.6 Hz, 2 H) 2.31 (t, J=7.3 Hz, 2 H) 1.60 - 1.73 (m, 7

H) 1.36 - 1.45 (m, 2 H) 0.95 (t, J=7.6 Hz, 3 H). 13C NMR (100 MHz, CHLOROFORM-d) δ

173.8, 173.5, 141.8, 130.5, 130.4, 129.0, 128.9, 127.2, 72.6, 62.2, 61.8, 36.2, 35.5, 34.4, 31.2,

28.9, 25.0, 18.6.

O

O

OH

OH

1,3-dihydroxypropan-2-yl 6-(biphenyl-4-yl)hexanoate, 35, AM9090. The reaction for 34 also

produced 35 (27 mg, 38%) as a white solid. Rf = 0.27 (70% ethyl acetate / hexanes). Mp 78-80

°C. 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.55 - 7.60 (m, 2 H) 7.49 - 7.53 (m, 2 H)

7.39 - 7.45 (m, 2 H) 7.30 - 7.35 (m, 1 H) 7.22 - 7.25 (m, 2 H) 4.92 (quin, J=4.8 Hz, 1 H) 3.81 (t,

J=5.1 Hz, 4 H) 2.66 (t, J=7.6 Hz, 2 H) 2.39 (t, J=7.6 Hz, 2 H) 2.08 (t, J=5.9 Hz, 2 H) 1.64 - 1.74

(m, 4 H) 1.37 - 1.45 (m, 2 H). 13C NMR (100 MHz, CHLOROFORM-d) δ 174.2, 141.8, 141.3,

138.9, 129.0, 128.9, 127.2, 72.6, 62.7, 35.5, 34.5, 31.2, 28.9, 25.1.

172

O

O

O

O

OH

(Z)-1-(butyryloxy)-3-hydroxypropan-2-yl 6-(biphenyl-4-yl)hex-5-enoate, 36, AM10321. The

procedure for 17 was followed to synthesize 36 (10 mg, 8%) as a white solid. Rf = 0.79 (75%


7.39 - 7.48 (m, 2 H) 7.31 - 7.37 (m, 3 H) 6.49 (d, J=11.7 Hz, 1 H) 5.66 (dt, J=11.7, 7.3 Hz, 1 H)

5.07 (quin, J=5.1 Hz, 1 H) 4.30 (dd, J=11.7, 4.4 Hz, 1 H) 4.21 (dd, J=11.7, 5.9 Hz, 1 H) 3.70 (d,

J=5.1 Hz, 2 H) 2.37 - 2.48 (m, 4 H) 2.28 (t, J=7.3 Hz, 2 H) 1.83 (dt, J=15.2, 7.4 Hz, 2 H) 1.62 (s,

2 H) 1.26 (t, J=7.3 Hz, 1 H) 0.93 (t, J=7.7 Hz, 3 H).

O

O

OH

OH

(Z)-1,3-dihydroxypropan-2-yl 6-(biphenyl-4-yl)hex-5-enoate, 37, AM10322. The reaction for

36 also produced 37 (64 mg, 65%) as a white solid. Rf = 0.29 (75% ethyl acetate / hexanes). 1H

NMR (500 MHz, CHLOROFORM-d) δ ppm 7.56 - 7.63 (m, 4 H) 7.42 - 7.47 (m, 2 H) 7.31 -

7.37 (m, 3 H) 6.50 (d, J=11.7 Hz, 1 H) 5.66 (dt, J=11.5, 7.2 Hz, 1 H) 4.89 (quin, J=4.8 Hz, 1 H)

3.77 (d, J=4.9 Hz, 4 H) 2.38 - 2.47 (m, 6 H) 1.83 (quin, J=7.4 Hz, 2 H). 13C NMR (100 MHz,

CHLOROFORM-d) δ 174.0, 141.0, 139.7, 136.7, 129.9, 129.6, 129.2, 127.4, 127.3, 126.9, 75.4,

62.5, 34.0, 28.2, 25.3.

173

O

OO

O

O

O

2-(pent-4-ynoyloxy)propane-1,3-diyl dibutyrate, 39. The procedure for 15 was followed to

synthesize 39 (3.27 g, 73%) as a colorless oil. Rf = 0.63 (35% ethyl acetate / hexanes). 1H NMR

(399 MHz, CHLOROFORM-d) δ ppm 5.27 - 5.34 (m, 1 H) 4.32 (dd, J=12.5, 3.7 Hz, 2 H) 4.17

(dd, J=11.7, 5.9 Hz, 2 H) 2.55 - 2.62 (m, 2 H) 2.48 - 2.55 (m, 2 H) 2.31 (t, J=7.3 Hz, 4 H) 1.98

(t, J=2.2 Hz, 1 H) 1.65 (sxt, J=7.5 Hz, 4 H) 0.91 - 0.99 (m, 6 H).

O

O

O

O

O

O

O

2-(5-(4-phenoxyphenyl)pent-4-ynoyloxy)propane-1,3-diyl dibutyrate, 41. The procedure for

27 was followed to synthesize 41 (800 mg, 42%) as a colorless oil. Rf = 0.56 (25% ethyl acetate /


H) 7.01 (d, J=7.3 Hz, 2 H) 6.87 - 6.93 (m, 2 H) 5.25 - 5.36 (m, 1 H) 4.33 (dd, J=12.1, 4.0 Hz, 2

H) 4.18 (dd, J=11.7, 5.9 Hz, 2 H) 2.69 - 2.76 (m, 2 H) 2.61 - 2.68 (m, 2 H) 2.24 - 2.31 (m, 4 H)

1.63 (sxt, J=7.3 Hz, 4 H) 0.89 - 0.96 (m, 6 H) 0.00 (s, 1 H). 13C NMR (100 MHz,

CHLOROFORM-d) δ 173.4, 171.3, 157.4, 156.8, 133.4, 130.1, 123.9, 119.5, 118.6, 118.3, 87.1,

81.0, 69.7, 62.2, 36.1, 33.8, 18.6, 15.5, 13.8.

O

O

O

OH

OH

174

1,3-dihydroxypropan-2-yl 5-(4-phenoxyphenyl)pent-4-ynoate, 42, AM10330. The procedure

for 31 was followed to synthesize 42 (51 mg, 71%) as a colorless oil. Rf = 0.24 (75% ethyl

acetate / hexanes). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 7.32 - 7.38 (m, 4 H) 7.09 -

7.16 (m, 1 H) 6.98 - 7.04 (m, 2 H) 6.88 - 6.93 (m, 2 H) 5.00 (quin, J=4.8 Hz, 1 H) 3.82 - 3.89 (m,

4 H) 2.73 - 2.79 (m, 2 H) 2.67 - 2.73 (m, 2 H) 1.99 - 2.09 (m, 2 H).

O

O

O

O

O

O

O

(Z)-2-(5-(4-phenoxyphenyl)pent-4-enoyloxy)propane-1,3-diyl dibutyrate, 43. The procedure

for 33 was followed to synthesize 43 (222 mg, 63%) as a colorless oil. Rf = 0.60 (25% ethyl


7.26 (m, 2 H) 7.08 - 7.14 (m, 1 H) 7.03 (d, J=8.8 Hz, 2 H) 6.95 - 7.00 (m, 2 H) 6.43 (d, J=11.7

Hz, 1 H) 5.57 (dt, J=11.5, 7.1 Hz, 1 H) 5.21 - 5.35 (m, 1 H) 4.31 (dd, J=12.1, 4.0 Hz, 2 H) 4.15

(dd, J=11.7, 5.9 Hz, 1 H) 2.62 - 2.71 (m, 2 H) 2.44 - 2.51 (m, 2 H) 2.29 (t, J=7.3 Hz, 4 H) 1.59 -

1.69 (m, 4 H) 0.94 (t, J=7.7 Hz, 6 H). 13C NMR (100 MHz, CHLOROFORM-d) δ 173.4, 172.3,

156.3, 132.4, 130.4, 130.0, 123.6, 119.2, 118.7, 94.7, 72.7, 62.2, 36.1, 34.5, 18.6, 13.9, 12.8.

O

O

OH

OH

O

(Z)-1,3-dihydroxypropan-2-yl 5-(4-phenoxyphenyl)pent-4-enoate, 44, AM10331. The



7.22 - 7.28 (m, 2 H) 7.08 - 7.14 (m, 1 H) 7.00 - 7.06 (m, 2 H) 6.95 - 7.00 (m, 2 H) 6.44 (d,

J=11.7 Hz, 1 H) 5.58 (dt, J=11.7, 7.0 Hz, 1 H) 4.92 (quin, J=4.8 Hz, 1 H) 3.75 - 3.85 (m, 4 H)

175

2.69 (q, J=7.3 Hz, 2 H) 2.53 (t, J=7.3 Hz, 2 H) 2.24 - 2.32 (m, 2 H). 13C NMR (100 MHz,

CHLOROFORM-d) δ 173.3, 157.2, 156.4, 130.4, 130.0, 129.9, 129.6, 123.6, 119.2, 118.7, 75.4,

62.5, 34.7, 24.3.

O O

Br

3-bromopentane-2,4-dione, 46. Bromine (5.4 mL, 105 mmol) was slowly added drop wise to a

solution of acetyl acetone (10.0 g, 100 mmol) in 40 mL of a 1:1 solution water:carbon

tetrachloride. After completion the reaction was diluted with DCM, and the organic layer was

washed with water (3x) and sat. sodium thiosulfate solution (2x). The organic layer was dried

over MgSO4, and evaporation of solvent under reduced pressure gave 3-bromopentane-2,4-dione

(11.3 g, 63%) as an oil. 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 15.86 (s, 1 H) 4.74 (s, 1

H) 2.61 (s, 3 H) 2.42 (s, 3 H) 2.34 - 2.35 (m, 6 H).

O

OO

O

(5Z,8Z,11Z,14Z)-2,4-dioxopentan-3-yl icosa-5,8,11,14-tetraenoate, 47, AM10332.

Arachidonic acid (300 mg, 1.0 mmol) was added to a solution of 60% NaH in mineral oil (40

mg, 1.0 mmol) in 5 mL of anhydrous DMF at 0 °C. After 0.5 h, 46 (35 mg, 0.2 mmol) was added

to the solution and was heated to 50 °C for 2 h. Upon completion the reaction was diluted with

water and ether. The organic layer was separated, dried over MgSO4, concentrated, and

chromatographed on silica gel to yield, 47 (64 mg, 79%) as a colorless oil. Rf = 0.71 (35% ethyl


2.88 (m, 6 H) 2.49 - 2.55 (m, 2 H) 2.30 (s, 4 H) 2.14 - 2.22 (m, 2 H) 2.06 (q, J=6.9 Hz, 2 H) 2.01

(s, 2 H) 1.75-1.83 (m, 2 H) 1.32 - 1.40 (m, 2 H) 1.24 - 1.32 (m, 4 H) 0.89 (t, J=7.0 Hz, 3 H). 13C

176

NMR (100 MHz, CHLOROFORM-d) δ 173.8, 170.8, 169.7, 130.7, 129.4, 128.8, 128.6, 128.5,

128.3, 128.1, 127.7, 85.2, 45.7, 33.3, 31.7, 29.6, 27.6, 27.5, 26.6, 25.9, 24.9, 24.8, 22.8, 14.3.

O

OOH

OH

(5Z,8Z,11Z,14Z)-((2S,4S)-2,4-dihydroxypentan-3-yl) icosa-5,8,11,14-tetraenoate, ±48,

AM10336. A solution of 47 (607 mg, 1.51 mmol) in 5 mL of THF was added dropwise to a

suspension of NaBH4 (58mg, 1.51mmol) in 5 mL of MeOH at -78 °C. After 2h, 0.1M HCl was

slowly added to the reaction mixture which was then diluted with H2O and Et2O. The ethereal

layer was separated, dried over MgSO4, and evaporated under reduced pressure. The oily

residue was chromatographed on silica gel to yield ±48, (255 mg, 42%) as a colorless oil. Rf =

0.42 (10% acetone / CH2Cl2). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.31 - 5.45 (m, 8

H) 4.67 (dd, J=5.2, 2.7 Hz, 1 H) 4.23 (quind, J=6.3, 6.3, 6.3, 6.3, 2.4 Hz, 1 H) 4.04 - 4.12 (m, 1

H) 2.78 - 2.87 (m, 6 H) 2.62 (dd, J=5.5, 2.4 Hz, 2 H) 2.42 (t, J=7.3 Hz, 2 H) 2.14 (dt, J=7.9, 6.7

Hz, 2 H) 2.06 (q, J=7.3 Hz, 2 H) 1.75 (quin, J=7.5 Hz, 2 H) 1.33 - 1.39 (m, 2 H) 1.27 - 1.33 (m,

4 H) 1.25 (d, J=6.7 Hz, 3 H) 1.19 (d, J=6.7 Hz, 3 H) 0.89 (t, J=6.7 Hz, 2 H). 13C NMR (100

MHz, CHLOROFORM-d) δ 173.7, 130.9, 129.4, 129.1, 128.9, 128.6, 128.4, 128.1, 127.8, 78.6,

68.6, 66.8, 34.0, 31.9, 29.7, 27.6, 26.9, 26.0, 25.2, 22.9, 19.9, 19.8, 14.4.

O

OOH

OH

(5Z,8Z,11Z,14Z)-((2R,3s,4S)-2,4-dihydroxypentan-3-yl) icosa-5,8,11,14-tetraenoate, 49,

AM10335. 1.0M Et2BOMe in THF (0.5 ml, 0.5 mmol) was added to a solution of 47 (200 mg,

177

0.5 mmol) in 6 mL of a 5:1 THF/methanol solution cooled to -78 °C. After 20 minutes NaBH4

was added portion wise (38 mg, 1.0 mmol). After 2 h the reaction was warmed to rt and then

quenched with 1M HCl. The solution was then diluted with H2O and Et2O. The ethereal layer

was separated, dried over MgSO4, and evaporated under reduced pressure. The oily residue was

chromatographed on silica gel to yield 49 (27 mg, 13%) as a colorless oil. Rf = 0.35 (10%

acetone / CH2Cl2). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.29 - 5.46 (m, 8 H) 4.68 (t,

J=6.4 Hz, 1 H) 3.98 - 4.08 (m, 2 H) 2.76 - 2.88 (m, 6 H) 2.42 (d, J=3.1 Hz, 2 H) 2.37 (t, J=7.3

Hz, 2 H) 2.13 (q, J=7.3 Hz, 2 H) 2.06 (q, J=7.3 Hz, 2 H) 1.73 (quin, J=7.5 Hz, 2 H) 1.25 - 1.40

(m, 6 H) 1.22 (d, J=6.7 Hz, 6 H) 0.87 - 0.91 (m, 3 H). 13C NMR (126 MHz, CHLOROFORM-d)

δ 173.7, 130.8, 129.3, 129.0, 128.9, 128.5, 128.3, 128.1, 127.8, 80.4, 69.1, 34.0, 31.8, 29.6, 27.5,

26.8, 25.9, 25.1, 22.8, 19.9, 14.3.

O

OOH

OH

(5Z,8Z,11Z,14Z)-((2R,3r,4S)-2,4-dihydroxypentan-3-yl) icosa-5,8,11,14-tetraenoate, 50

AM10334. The reaction for 49 also produced 50 (69 mg, 35%) as a colorless oil. Rf = 0.25 (10%

acetone / CH2Cl2). 1H NMR (500 MHz, CHLOROFORM-d) δ ppm 5.30 - 5.46 (m, 8 H) 4.73 (t,

J=3.1 Hz, 2 H) 4.13 (quind, J=6.3, 6.3, 6.3, 6.3, 3.1 Hz, 2 H) 2.79 - 2.87 (m, 7 H) 2.47 (t, J=7.3

Hz, 2 H) 2.36 - 2.40 (m, 2 H) 2.13 - 2.19 (m, 2 H) 2.06 (q, J=7.3 Hz, 2 H) 1.77 (quin, J=7.5 Hz,

2 H) 1.33 - 1.40 (m, 2 H) 1.27 - 1.33 (m, 4 H) 1.20 (d, J=6.7 Hz, 6 H) 0.89 (t, J=7.0 Hz, 3 H). 13C

NMR (126 MHz, CHLOROFORM-d) δ 173.7, 130.9, 129.3, 129.1, 128.9, 128.5, 128.3, 128.1,

127.8, 80.6, 69.1, 34.1, 31.8, 29.6, 27.5, 26.8, 25.9, 25.1, 22.8, 19.9, 14.4.

O

O

178

1-(oxiran-2-yl)ethanone, 52. Hydrogen peroxide (30%, 1.2 mL, 10.7 mmol) was added to a

stirred solution of methyl vinyl ketone 51 (0.593 mL, 7.1 mmol) and sodium bicarbonate (5.9 g,

71 mmol) in water (20 mL). After 3 h the product was extracted with ether (3x) and the organic

layer was then concentrated. The resulting residue was chromatographed on silica gel to yield 52

(70 mg, 11%) as an oil. Rf = 0.23 (10% ethyl acetate / hexanes). 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 3.41 (dd, J=4.3, 2.4 Hz, 1 H) 2.99 - 3.05 (m, 1 H) 2.90 (dd, J=5.5,

2.4 Hz, 1 H) 2.07 (s, 3 H).

O

O

OH

OH

(5Z,8Z,11Z,14Z)-1,3-dihydroxybutan-2-yl icosa-5,8,11,14-tetraenoate, 53. Immobillized

Candida antarctica (200 mg) was added to a stirred solution of 57 (100 mg, 0.1 mmol) in

anhydrous ethanol (1.5 mL) at room temperature. The reaction was stirred for 2 h, at which an

additional lipase (100 mg) was added. After 2 h the reaction mixture was filtered, the ethanol

was removed under reduced pressure and the residue was chromatographed on a silica gel to

yield AM10342 (29 mg, 73%) as a colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ ppm

5.31 - 5.46 (m, 8 H) 4.77 (td, J=4.9, 3.7 Hz, 1 H) 4.07 (qd, J=6.7, 1.8 Hz, 1 H) 3.92 (dd, J=12.2,

4.9 Hz, 1 H) 3.84 (dd, J=12.2, 3.7 Hz, 1 H) 2.77 - 2.88 (m, 6 H) 2.40 (t, J=7.3 Hz, 2 H) 2.14 (q,

J=6.7 Hz, 2 H) 2.06 (q, J=6.9 Hz, 2 H) 1.74 (quin, J=7.3 Hz, 2 H) 1.33 - 1.40 (m, 2 H) 1.27 -

1.33 (m, 4 H) 1.26 (d, J=6.1 Hz, 4 H) 0.89 (t, J=6.7 Hz, 3 H).

O

O

O

O

O

C19H31

O

C19H31

179

1,2,3-tri[(5Z,8Z,11Z,14Z)-icosa-5,8,11,14-tetraenoate]-4-methyl-sn-glycerol, 57. EDCI

(1.33g, 6.9 mmol), DMAP (56 mg, 0.46 mmol), and 1,3-dibutoylglycerol (700 mg, 3.0 mmol)

were added to a stirred solution of arachidonic acid (700 mg, 2.31 mmol) in anhydrous CH2Cl2

(15 mL) at 0°C. The reaction was allowed to stir under argon for 4 hours. Upon completion the

reaction was diluted with CH2Cl2 and water. The organic layer was concentrated and the residue

was chromatographed on silica gel to yield 57 (1.15 g, 96%) as an oil. 1H NMR (500 MHz,

CHLOROFORM-d) δ ppm 5.29 - 5.44 (m, 24 H) 5.14 - 5.19 (m, 1 H) 5.07 - 5.14 (m, 1 H) 4.31

(dd, J=11.9, 3.4 Hz, 1 H) 4.13 (dd, J=11.9, 7.0 Hz, 1 H) 2.75 - 2.88 (m, 18 H) 2.27 - 2.36 (m, 6

H) 2.08 - 2.15 (m, 6 H) 2.05 (q, J=7.3 Hz, 6 H) 1.64 - 1.74 (m, 6 H) 1.26 - 1.40 (m, 18 H) 1.25

(d, J=6.1 Hz, 3 H) 0.89 (t, J=7.0 Hz, 9 H).

180

4.9 References 1. Katona, I.; Sperlágh, B.; Sık, A.; Käfalvi, A.; Vizi, E. S.; Mackie, K.; Freund, T. F. Presynaptically Located CB1 Cannabinoid Receptors Regulate GABA Release from Axon Terminals of Specific Hippocampal Interneurons. The Journal of Neuroscience 1999, 19, 4544-4558. 2. Sugiura, T.; Kodaka, T.; Nakane, S.; Miyashita, T.; Kondo, S.; Suhara, Y.; Takayama, H.; Waku, K.; Seki, C.; Baba, N.; Ishima, Y. Evidence That the Cannabinoid CB1 Receptor Is a 2-Arachidonoylglycerol Receptor. Journal of Biological Chemistry 1999, 274, 2794-2801. 3. Marsicano, G.; Lutz, B. Expression of the cannabinoid receptor CB1 in distinct neuronal subpopulations in the adult mouse forebrain. European Journal of Neuroscience 1999, 11, 4213-4225. 4. Abadji, V.; Lin, S.; Taha, G.; Griffin, G.; Stevenson, L. A.; Pertwee, R. G.; Makriyannis, A. (R)-Methanandamide: A Chiral Novel Anandamide Possessing Higher Potency and Metabolic Stability. Journal of Medicinal Chemistry 1994, 37, 1889-1893. 5. Parkkari, T.; Myllymäki, M.; Savinainen, J. R.; Saario, S. M.; Castillo-Meléndez, J. A.; Laitinen, J. T.; Nevalainen, T.; Koskinen, A. M. P.; Järvinen, T. α-Methylated derivatives of 2-arachidonoyl glycerol: Synthesis, CB1 receptor activity, and enzymatic stability. Bioorganic & Medicinal Chemistry Letters 2006, 16, 2437-2440. 6. Parkkari, T.; Salo, O. M. H.; Huttunen, K. M.; Savinainen, J. R.; Laitinen, J. T.; Poso, A.; Nevalainen, T.; Järvinen, T. Synthesis and CB1 receptor activities of dimethylheptyl derivatives of 2-arachidonoyl glycerol (2-AG) and 2-arachidonyl glyceryl ether (2-AGE). Bioorganic & Medicinal Chemistry 2006, 14, 2850-2858. 7. Suhara, Y.; Oka, S.; Kittaka, A.; Takayama, H.; Waku, K.; Sugiura, T. Synthesis and biological evaluation of several structural analogs of 2-arachidonoylglycerol, an endogenous cannabinoid receptor ligand. Bioorganic & Medicinal Chemistry 2007, 15, 854-867. 8. Vadivel, S. K.; Vardarajan, S.; Duclos Jr, R. I.; Wood, J. T.; Guo, J.; Makriyannis, A. Conformationally constrained analogues of 2-arachidonoylglycerol. Bioorganic & Medicinal Chemistry Letters 2007, 17, 5959-5963. 9. Brizzi, A.; Cascio, M. G.; Frosini, M.; Ligresti, A.; Aiello, F.; Biotti, I.; Brizzi, V.; Pertwee, R. G.; Corelli, F.; Di Marzo, V. Resorcinol-sn-Glycerol Derivatives: Novel 2-Arachidonoylglycerol Mimetics Endowed with High Affinity and Selectivity for Cannabinoid Type 1 Receptor. Journal of Medicinal Chemistry 2011. 10. Wagner, J. A.; Járai, Z.; Bátkai, S.; Kunos, G. Hemodynamic effects of cannabinoids: coronary and cerebral vasodilation mediated by cannabinoid CB1 receptors. European Journal of Pharmacology 2001, 423, 203-210. 11. Boswinkel, G.; Derksen, J.; van't Riet, K.; Cuperus, F. Kinetics of acyl migration in monoglycerides and dependence on acyl chainlength. Journal of the American Oil Chemists' Society 1996, 73, 707-711. 12. Szabo, B.; Nordheim, U.; Niederhoffer, N. Effects of Cannabinoids on Sympathetic and Parasympathetic Neuroeffector Transmission in the Rabbit Heart. Journal of Pharmacology and Experimental Therapeutics 2001, 297, 819-826. 13. Fowler, C. J. Anandamide uptake explained? Trends in Pharmacological Sciences 2012, 33, 181-185. 14. Stelt, M. v. d.; Kuik, J. A. v.; Bari, M.; Zadelhoff, G. v.; Leeflang, B. R.; Veldink, G. A.; Finazzi-Agro`, A.; Vliegenthart, J. F. G.; Maccarrone, M. Oxygenated Metabolites of

181

Anandamide and 2-Arachidonoylglycerol: Conformational Analysis and Interaction with Cannabinoid Receptors, Membrane Transporter, and Fatty Acid Amide Hydrolase. Journal of Medicinal Chemistry 2002, 45, 3709-3720. 15. Mechoulam, R.; Ben-Shabat, S.; Hanus, L.; Ligumsky, M.; Kaminski, N. E.; Schatz, A. R.; Gopher, A.; Almog, S.; Martin, B. R.; Compton, D. R.; Pertwee, R. G.; Griffin, G.; Bayewitch, M.; Barg, J.; Vogel, Z. Identification of an endogenous 2-monoglyceride, present in canine gut, that binds to cannabinoid receptors. Biochemical Pharmacology 1995, 50, 83-90. 16. Ghafouri, N.; Tiger, G.; Razdan, R. K.; Mahadevan, A.; Pertwee, R. G.; Martin, B. R.; Fowler, C. J. Inhibition of monoacylglycerol lipase and fatty acid amide hydrolase by analogues of 2-arachidonoylglycerol. British Journal of Pharmacology 2004, 143, 774-784. 17. Sugiura, T.; Kondo, S.; Sukagawa, A.; Nakane, S.; Shinoda, A.; Itoh, K.; Yamashita, A.; Waku, K. 2-Arachidonoylgylcerol: A Possible Endogenous Cannabinoid Receptor Ligand in Brain. Biochemical and Biophysical Research Communications 1995, 215, 89-97. 18. Mukherjee, S.; Adams, M.; Whiteaker, K.; Daza, A.; Kage, K.; Cassar, S.; Meyer, M.; Yao, B. B. Species comparison and pharmacological characterization of rat and human CB2 cannabinoid receptors. European Journal of Pharmacology 2004, 505, 1-9. 19. Goparaju, S. K.; Ueda, N.; Yamaguchi, H.; Yamamoto, S. Anandamide amidohydrolase reacting with 2-arachidonoylglycerol, another cannabinoid receptor ligand. FEBS Letters 1998, 422, 69-73. 20. Croci, T.; Manara, L.; Aureggi, G.; Guagnini, F.; Rinaldi-Carmona, M.; Maffrand, J.-P.; Le Fur, G.; Mukenge, S.; Ferla, G. In vitro functional evidence of neuronal cannabinoid CB1 receptors in human ileum. British Journal of Pharmacology 1998, 125, 1393-1395. 21. Rodrıguez, J. J.; Mackie, K.; Pickel, V. M. Ultrastructural Localization of the CB1 Cannabinoid Receptor in μ-Opioid Receptor Patches of the Rat Caudate Putamen Nucleus. The Journal of Neuroscience 2001, 21, 823-833.

182

CHAPTER 5

N-ACYLETHANOLAMINE-HYDROLYZING ACID AMIDASE INHIBITORS

183

5.1 Introduction and Background

NAEs are signaling molecules consisting of a fatty acid coupled to ethanolamine through

an amide bond and are found in plants and animals.1, 2 Anandamide has been the most studied

NAE because of its role in the endocannabinoid system.3 Palmitoylethanolamine (PEA) and

Oleoylethanolamine (OEA) are NAEs that have been extensively studied and which exhibit anti-

inflammatory properties through their activation of the PPARα receptor,4 in addition to

producing effects relating to neuroprotection,5 and analgesia.6 PEA also has an affinity to the

GPR55 and GPR119 receptors;7 and whether PEA activates the cannabinoid receptors (CB1 &

CB2) has been debated.8 It has been postulated that PEA activates the CB2 receptor but it may be

that the effects of PEA are not directly related to receptor activation. This may be the result of

an ‘entourage effect’ where PEA and other NAEs compete for their degradation by FAAH, and

thus increase the biological activity of anandamide by impeding its degradation.9 Studies

utilizing FAAH-deficient mice10-12 and selective FAAH inhibitors13, 14 suggest that FAAH is

primarily responsible for the metabolism of NAEs in the mammalian brain, while NAAA

metabolizes PEA in many mammalian tissues, organs, and some components of the immune

system.15-18 Thus, inhibition of NAAA and the subsequent increase of PEA could amplify anti-

inflammatory and anti-nociceptive effects of greater anandamide concentrations, as well as

increased PEA levels.

NAEs were believed to be degraded only by FAAH to the free fatty acid and

ethanolamine, until recently when NAAA was purified and characterized to be distinctively

different than FAAH and responsible for NAE metabolism.19 NAAA is primarily localized in

lysosomes,20 while FAAH is distributed in the cytosolic and luminal sides of intracellular

membranes. NAAA also has a modest homology (30%) with FAAH which was determined after

184

the enzyme was cloned from rat, mouse, and human cDNA.21 NAAA belongs to the N-terminal

nuecleophile hydrolase superfamily, containing an N-terminal cysteine (126 in human) as the

catalytic residue.21, 22 Most importantly, NAAA has been shown to have optimum catalytic

activity at an acidic pH (4.5) compared to FAAH which has an ideal activity at pH 8.5-10.23 The

primary substrates of each enzyme differ as well. NAAA hydrolyzes PEA 40 times greater than

that of anandamide,16 while FAAH hydrolyzed anandamide eight times greater than PEA.24

5.1.1 Current NAAA inhibitors

After the identification,17 cloning, and characterization of NAAA,16 several research

efforts began into the development of novel inhibitors to increase NAE levels and thus amplify

their pharmacological benefits. The first reported NAAA inhibitors were a series of esters,

retroesters, and retroamides of palmitic acid (Figure 5.1).25 These compounds were assayed

utilizing a radioactive TLC assay, with NAAA from rat lung, at a concentration of 100 μM for

each inhibitor. Analogs of palmitoyl ester 1, reteroester 2 and the retroamine 3, had an inhibititon

of 84%, 71% and 77% respectively. PEA, the natural substrate for NAAA, had an inhibition of

34% at a concentration of 100 μM. While the first reported inhibitors were not a great

improvements over the substrate for inhibiting NAAA, compounds 2 and 3 showed that the

sensitivity of NAAA to retroesters and retroamides was different to that for FAAH, as no

significant inhibition of FAAH at the same concentration by 2 and 3 was observed.25 Saturnino,

et al., completed an SAR study based on these esters, retroester and retroamides using the human

NAAA enzyme. From this study, 5 exhibited an improved inhibition of 85% at 50 μM.26

185

O

O1

O

O

2

NH

O

HO

3

O

NH

OH

4

O

O5

Figure 5.1 Best NAAA inhibitors derived from palmitic esters, 1 & 5; retroesters, 2;

retroamides, 3; along with palmitoylethanolamine (PEA) 4.

More recently, a new class of inhibitors containing a β-lactone displayed an improved

inhibition of NAAA. Using computer models based on the conserved catalytic N-terminal

region of conjugated bile acid hydrolase, compound 6 (Figure 5.2) was designed, synthesized

and shown to have an IC50 of 420 nM with rat NAAA. It also appears that NAAA displays

selective recognition, as the enantiomer of 6, compound 7, had a dramatic reduction of inhibition

to 6000 nM. In vivo experiments using 6 to inhibit NAAA increased endogenous levels of PEA,

thus preventing acute inflammation which appears to act through PPAR-α.27

This β-lactone scaffold led to the development of potent inhibitors 8 (102 nM), 9 (90

nM), and 10 (115 nM).28 Duranti, et al., continued SAR studies of this scaffold as the amide-

lactone functionality lacks the desired chemical stability, as these α-amino-β-lactones react with

other bionucleophiles and can be hydrolyzed in aqueous media. The chemical stability of these

compounds was improved by replacing the amide with a carbamate moiety, resulting in

compound 11 (Figure 5.2), with an IC50 of 130 nM, and a 10 fold increase in chemical stability

in rat and human plasma, and in the presence of bovine serum albumin.29

186

O

NHO

O O

NHO

O

6 7

O

NHO

O O

NHO

O

O

O

NHO

O

8 9 10

O

NHO

O

O

11

Figure 5.2 Reported β-lactone NAAA inhibitors

5.2 Design, Synthesis, and Biological Evaluation of NAAA Inhibitors

5.2.1 Retroamides

In our search to find a class of compounds that selectively inhibited NAAA over other

enzymes present in the endocannabinoid system, we decided to start with a series of retroamides.

This decision was based on that the observation that retroamides were known to be more stable

in the presence of FAAH and MGL, and would hopefully have selectivity towards NAAA.30 All

retroamides (Table 5.1) were prepared starting with palmitylamine. The formation of the amide

187

bond was achieved with either an acyl chloride in the presence of catalytic DMAP (12-16 and

18, Scheme 5.1), or an acid using EDCI and DMAP (17).

Compound 20 was synthesized from the p-nitrophenyl derivative of 18 which was treated with

palladium and hydrogen gas to yield amine 19. 19 was then treated with 1,1-thiocarbonyl-2(1H)-

dipyridone to yield isothiocyanate 20. While each of these retroamides did not show significant

activity at either FAAH or MGL (Table 5.1), there was also no significant inhibition of NAAA.

It appears that aryl and large cyclic groups placed at the head position of the compound were not

suitable for inhibition of NAAA, unlike the low μM inhibition observed from compounds 1 and

5.

Scheme 5.1 Synthesis of retroamide inhibitors

H2NO

ClR

NH

R

Ocat. DMAPCH2Cl2, 0 °C

12-16, 18

H2NO

OH

NH

OEDCI, DMAPCH2Cl2, 0 °C

17

1878%

Pd/C, EtOAcNH

O

H2N54%

NH

O

SCN

NO

NS O

14

19 20

188

Table 5.1 IC50 values for compounds 12-20 towards NAAA, FAAH, and MGL enzymes. Cmpd

No. AM Structure IC50 (μM) hNAAA rFAAH hMGL

12 9024 NH

O

~100 >100 >100

13 9026 NH

O

O 10-100 - -

14 9027 NH

O

~100 >100 ~100

15 9028 NH

O

~100 >100 ~100

16 9030 NH

O

F >100 >100 >100

17 9031 O

HN

~100 >100 >100

18 9033 O

NH

O2N ~100 >100 >100

19 9034 O

NH

H2N >100 >100 >100

20 9035 O

NH

SCN >100 >100 >100

5.2.2 Carbonates and Carbamates

To capitalize on the catalytic cysteine residue of NAAA, we further investigated the

activity of carbonate and carbamate containing ligands as possible inhibitors. To maximize the

electrophilic nature of the functional group, a p-nitrobenzene was introduced to act as a good

leaving group. The electron withdrawing properties of the nitro group makes the phenyl a better

leaving group after nucleophilic attack of the electrophilic carbonyl moiety.

The synthesis of carbonates was conducted by reacting a series of alcohols 21-23 with p-

nitrophenyl chloroformate 24, in the presence of triethylamine and DMAP (Scheme 5.2).

Carbamates 33-36 were synthesized with a p-cyanophenyl group as the leaving group. While

cyano is a weaker electron withdrawing group than nitro, it provides more functional stability.

189

These were synthesized starting with the treatment of alkylamines 28-31 with triethylamine and

phosgene to form the intermediate carbamic chloride, which was subsequently treated with 4-

hydroxybenzonitrile 32 to yield carbamates 33-36 (Scheme 5.3).

10-phenyldecanol 22 was converted to azide 37 with DPPA and DBU in excellent yield.

In two steps azide 37 was reduced to the amine and reacted with 24 to yield carbamate 38 in low

yield. Amines 39 and 40 were reacted with 24 in the presence of triethylamine and DMAP to

yield carbamates 41 and 42 in 98% and 32% yields respectively (Scheme 5.3).

SAR studies noted the differences of the IC50 values compared among enzymes NAAA,

FAAH and MGL (Table 5.2). Overall, carbamates exhibited greater NAAA inhibition as

compared to carbonates. The moderately potent carbonate 27 (AM9056) exhibited an IC50 of 5.8

μM towards NAAA, while having no inhibition of FAAH or MGL (>100 μM), however, it was

quickly apparent that the carbamates were generally more potent. Evaluation of SAR of

carbonates was not pursued at this time, although they may be selective.

Scheme 5.2 Synthesis of Carbonates 25-27

NO2

OCl

OOH

Xn

21 n = 15 X = CH322 n = 7 X = Ph23 n = 9 X = Ph

OX

nO

ONO2

25 n = 15 X = CH326 n = 7 X = Ph27 n = 9 X = Ph

24

Et3N, DMAPTHF

71-98%

190

Scheme 5.3 Synthesis of Carbamates 33-36, 38 and 40-41

NH2nn = 10, 12-14

28-31

CN

HO NHn

O

OCN

n = 10, 12-1433-36

NHn

O

ONO2

X NH2n

28 n = 11 X = CH339 n = 0 X = p-benzyloxybenzene

X

40 n = 11 X = CH341 n = 0 X = p-benzyloxybenzene

Et3N, DMAPTHF

23DPPA, DBUDMF, 120 °C

N3n

1) Pd/C H22) 24, CH2Cl2

37

32

NHn

O

ONO2

38

Et3N, COCl2CH2Cl2, 0 °C

n = 10 n = 10

12-79%

13%95%

32-98%24

With regards to carbamate inhibitors, the functional group para to the carbamate had a

slight effect with activity. Compounds 33 and 34 which had a cyano in the para position had a

higher IC50 (486 nM and 308 nM respectively) after 3h incubation as compared to nitro

carbamate 41 (IC50 = 255 nM). The biggest difference between these compounds was their

selectivity towards FAAH. Compound 33 was actually more selective towards FAAH (IC50 =

313 nM for FAAH, 486 nM for NAAA), while 34 had 2x the selectivity for NAAA (IC50 = 812

nM for FAAH, 308 nM for NAAA).

The SAR study of carbamates culminated with the introduction of shorter alkyl chain and

a phenyl group at the tail end. Compound 38 (AM9058) is our the most potent and selective

(22x) carbamate inhibitor with an IC50 of 32 nM and 716 nM for NAAA and FAAH respectively.

191

Replacement of the phenylalkyl chain with a more bulky two-ring system, 41, resulted in a drop

of inhibition towards NAAA.

These compounds were expected to behave as covalent inhibitors, which are indicated by

the large increase in inhibition when compounds are incubated with the enzyme for longer

periods of time. 33, 34 and 38 all showed improved inhibition when the incubation time was

increased from 15 m to 3 h, which allowed more covalent interactions to occur with the catalytic

cysteine of the NAAA enzyme before its activity to hydrolyze PAMCA is measured.

Table 5.2 Carbonates and Carbamates

R X( )n

O

O

Y

Cmpd No. AM R X Y N

IC50 (μM) NAAA NAAA

rFAAHb hMGLb 15m 3h

25 9048 CH3 O NO2 16 10-100 - >100 >100

26 9057 Ph O NO2 8 10-100 - 10-100 >100

27 9056 Ph O NO2 10 5.8±3.6a - >100 >100

33 9052 CH3 NH CN 11 1.3±0.16a 0.49±0.41a 0.31 ~10

34 9051 CH3 NH CN 13 1.4±0.11a 0.31±0.18a 0.81 1-10

35 9050 CH3 NH CN 14 ~10 - 1.9 >100

36 9049 CH3 NH CN 15 10-100 - ~10 >100

38 9058 Ph NH NO2 10 0.085±0.008a 0.032±0.004a 0.72 ~10

40 9054 CH3 NH NO2 11 0.26±0.012a 2.4 10-100

41 9072 O NH NO2 0 2.36 >100 >100

aData from three experiments each run in triplicate. bData from one experiment run in triplicate.

192

5.2.3 Optimization of AM9058

Of the carbonates and carbamates, AM9058 (38, Table 5.2) exhibited the greatest

inhibition of NAAA (IC50 = 31.9 nM) and was 22 times more selective over FAAH. In an

attempt to improve enzyme selectivity, various leaving groups were examined. This was

completed through parallel synthesis of 23 using a series of chloroformates 42-48 to yield

carbamates 49-55 in 25-100% yield (Scheme 5.4).

Scheme 5.4 Parallel synthesis of carbamates

23OCl

OR

42 R = Ph 46 = 2-NO2 43 R = 4-F 47 = 4-OMe44 R = 2-Cl 48 = 4-Br45 R = 3-CF3

DMAP, CH2Cl20 °C

25-100% ONH

OR

49 R = Ph 53 = 2-NO2 50 R = 4-F 54 = 4-OMe51 R = 2-Cl 55 = 4-Br52 R = 3-CF3

n n = 10

While it was hoped that improve the NAAA/FAAH selectivity of the carbamates would

be improved by altering the leaving group, the opposite effect was observed. Only 52 and 53

exhibited significant NAAA inhibition. Compound 52 had an IC50 of 7.35 μM for 15 m

preincubation which improved to 0.713 μM after 3 h preincubation. While 53 exhibited an IC50

of 4.08 μM after 15 m preincubation and an improved inhibition of 0.988 μM after 3 h

preincubation (Table 5.3). While these ligands exhibited no significant MGL inhibition; they all

were selective for FAAH over NAAA with all ligands exhibiting <μM (IC50) FAAH inhibition.

Compounds 49-51 and 54-55 all had IC50 values <100 nM towards FAAH, with 55 being the

most potent at 5.9 nM. It appears that the carbamate moiety may not be suitable for optimal

NAAA inhibitors, as they are prone to attack from the catalytic serine of FAAH.

193

Table 5.3 Carbamate inhibitors with alternate phenyl leaving groups

HN

10O

OR

Cmpd No. AM R

IC50 (μM) NAAA NAAA rFAAHb hMGLb 15m 3h

49 9061

10-100 - 0.02 10-100

50 9062 F

10-100 - 0.02 10-100

51 9063 Cl

10-100 - 0.08 ~10

52 9064

CF3

7.4±0.9a 0.71±0.04a 0.1 1-10

53 9065 NO2

4.1±0.9a 1.0±0.5a 0.40 >100

54 9066 O

>100 - 0.006 1-10

55 9067 Br

10-100 - 0.09 ~10

aData from three separate experiments run in triplicate. bData from one experiment run in triplicate.

5.2.4 Isothiocyanates

Based on cysteine being established as the catalytic amino acid residue of NAAA, and

also capable of inhibition through covalent interaction with carbamates and β-lactones, it was

hypothesized that isothiocyanates would behave as covalent inhibitors. This was based on

covalent interactions previously observed between the isothiocyanate moiety and cysteine

residues in the cannabinoid receptors.31-33 Also, isothiocyanates are not susceptible to

nucleophilic attack from serine, which should improve selectivity over FAAH.

194

Synthesis began with EDCI coupling of acids 56 and 90 with ethanolamine to yield

amides 57 and 91. The hydroxyl group of the ethanolamide was transformed to the azide using

DPPA and DBU, which was converted to the isothiocyanate with PPh3 and CS2 to yield

inhibitors 59 and 53 (Scheme 5.5).

Amines 60-65 and 94-101 were treated with 1,1`-thiocarbonyldipyridin-2(1H)-one to

yield isothicyanates 66-71 and 102-109 in excellent yields. This reagent is simpler and more

efficient compared to previous methods of using carbon disulfide and triethylamine followed by

treatement with p-toluenesulfonyl chloride. The reaction with 1,1`-thiocarbonyldipyridin-2(1H)-

one was carried out at room temperature and reaction went to completion within 30 m. Yields

were mostly quantitative

Phenylalkanols 72-74 and 22-23 were first converted to azides 75-78 and 37 which led to

inhibitors 79-83 through established conditions. Although the transformation of azides to

isothiocyanates with PPh3 and CS2 can take 48 h, this method is preferred to reducing the azide

to the amine first as loss of compound is observed during purification, even though the

subsequent step can be quantitative.

To synthesize 89, 2-bromonaphthalene 84 and methyl hept-6-ynoate were treated with

trans-dichlorobis(triphenylphosphine)palladium (II), copper iodide, and triethylamine

(Sonogashira conditions) in the microwave for 20 minutes at 100°C to yield the coupled product

85. The alkyne was reduced to alkane 86 with palladium and hydrogen. Reduction of methyl

ester 86 to the alcohol using lithium aluminumhydride yielded alcohol 87, which was converted

to isothiocyanate 89 utilizing existing methods.

195

Scheme 5.5 Synthesis of Isothiocyanate NAAA inhibitors

R OH

O

56 R = (CH2)6C10H790 R = C15H32

R NH

OOH

R NH

ONCS

R NH

ON3

57 R = (CH2)6C10H791 R = C15H32

58 R = (CH2)6C10H792 R = C15H32

59 R = (CH2)6C10H793 R = C15H32

R NH2 R NCS

66-71, 102-10960-65, 94-101

OHn

n = 4-6,8,10

N3n

n = 4-6,8,10

NCSn

n = 4-6,8,1079-8372-74, 22, 23 75-78, 37

4

Br

OH7

87

84 85

methyl hept-6-ynoate, Pd(PPh3)2Cl2, Et3N, CuI Pd/C, H2

O

O6

86O

O

N37

88

NCS7

89

PPh3, CS2, THFDPPA, DBU, DMF, 120 °C

LAH, THF

EDCI, DMAPethanolamine

DPPA, DBU, DMF, 120 °C

CS2, PPh3, THF

NO

NS O

DPPA, DBU, DMF, 120 °C CS2, PPh3, THF

Since NAAA hydrolyzes NAEs at the amide group, it was first decided to synthesize an

analog of PEA where the hydroxyl of the ethanolamide was replaced with an isothiocyanate

196

group. The hypothesis was that this amide side of the ligand interacts with the catalytic triad of

the enzyme, and by placing a group known for covalent interactions with cysteines near the site

of enzymatic attack, a preference will be had for the isothiocyanate and the compound will act as

an irreversible inhibitor. This was the case with 59 where an IC50 of 900 nM was observed

towards NAAA, while having no interaction with FAAH.

Compound 66 was a result of shortening of the alkyl chain to 15 carbons and removing

the amide group. This showed an improvement in inhibition to 350 nM (Table 5.4). Shortening

the chain further increased inhibition as seen with 67 (14 carbons, IC50 = 150 nM) and 68 (12

carbons, IC50 = 170 nM). A significant reduction in potency was observed when the chain was

shortened to 10 carbons (69) as the IC50 for inhibition was around 10 μM. A further decrease of

inhibition to >100 μM was seen when a methyl was introduced α to the isothiocyanate group

(70).

The length of ligand chain was further shortened and a phenyl group was also placed at

the tail to improve NAAA inhibition. Compounds 79-81 had alkyl spacers of four, five, and six

carbons respectively, although no significant inhibition was observed. When there were eight

carbons between the phenyl and isothiocyanate groups the IC50 improved to 410 nM (82). The

increase in chain length culminated with compound 83 (AM9053) which exhibited an IC50 of 39

nM towards the enzyme. This coincides with the length of the best alkyl chain based inhibitors

67 and 68 in terms of optimal length of the compound. To expand on AM9053, the phenyl was

replaced with the larger naphthyl group while shortening the chain to seven carbons in 89.

While an increase of inhibition was not observed, this compound was still one of the better

inhibitors, with an IC50 of 260 nM. Modification of 89 to include an amide group as in 93,

however, resulted in a drastic loss of inhibitory activity.

197

Based on results of the best ligand AM9053, compounds were made to produce a series

of ring systems in place of the alkyl tail. Compounds containing a combination of phenyl,

pyridinyl, piperazinyl, morpholinyl, piperidinyl, and/or pyrazinyl ring systems were examined.

Unfortunately, none of these ligands showed any inhibition towards NAAA.

Table 5.4 Inhibition data of isothiocyanate compoudns Cmpd

No. AM Structure IC50 (μM) hNAAA rFAAH hMGL

59 9019 O

NH

NCS14

0.90±0.07a ~100 -

66 9023 13NCS

0.35±0.07a 10-100 ~10

67 9046 12NCS

0.15±0.03a ~100 -

68 9045 10NCS

0.17±0.07a 10-100 -

69 9042 8NCS

~10 10-100 -

70 9043 8

NCS

>100 >100 -

71 9044 SNCS

O

>100 >100 -

79 9038 NCS4

>100 ~10 ~100

80 9037 NCS5

>100 10-100 ~100

81 9036 NCS6

>100 1-10 10-100

82 9047 NCS8

0.41±0.11a 10-100 ~100

83 9053 NCS10

0.039±0.009a ~100 >100

89 9060 NCS7

0.26±0.03a >100 ~100

198

93 9071 O

NH

NCS6

10-100 10-100 1-10

102 10318 NNCS

>100 - -

103 10323 NCS

NN

>100 - >100

104 10324 NCS

ON

>100 - >100

105 10325 NCS

N

>100 10-100 >100

106 10326 NCSO

N >100 >100 >100

107 10327 N

O

ONCS

10-100 10-100 ~10

108 10328 NCS

ON

N 10-100 10-100 10-100

109 10329 NCS

O

10-100 ~10 ~10

aAll data obtained from three separate experiments run in triplicate

5.3 Evaluation of AM9053 Mode of Inhibition

The mechanism of inactivation of NAAA by β-lactones has been recently reported to

proceed through an S-acylation by the catalytic N-terminal cysteine (Scheme 5.6).34 We have

recently corroborated this result as well as identifying irreversible inhibition through cysteine

attack on the urea moiety of AM6701 (Scheme 5.7).35

199

Scheme 5.6 S-alkylation inhibition mechanism of β-lactones

O

O NH

O

OREnzyme-S

SEnzyme

ONH

O

OR

HO

Scheme 5.7 Irreversible inhibition by AM6701

Enzyme-S NN

NN

O

N SEnzyme

O

N

Our most potent inhibitor AM9053 (83, Table 5.4) was expected to behave as an

irreversible inhibitor when the catalytic N-terminal cysteine attacks the electrophilic

isothiocyanate group to form a covalent bond.32, 33, 36

During NAAA inhibition assays of carbamate ligands, the amount of inhibition was

related to the preincubation time. Compounds 33, 34 and 38 (Table 5.2) were assayed with

preincubation times of 15 m and 3 h. A dramatic increase of inhibition was observed with

increased preincubation. This is consistent with a covalent interaction of between the carbamate

moiety and N-terminal catalytic cysteine of the enzyme. However, for concentration dependent

inhibition of NAAA by AM9053, when preincubated for either 15 m and 3 h, there was no

discernible difference in activity (Figure 5.3).

200

Figure 5.3 Concentration dependent inhibition of human NAAA by AM9053 15 minutes preincubation with the compound (filled circles, solid line) and after 3 hours preincubation (open circles, dotted line).

To confirm this observation, MALDI-TOF MS was utilized to examine a tryptic digest of

the catalytic region of NAAA with and without treatment of AM9053. The control (A, Figure

5.4) consisted of the tryptic digest of human NAAA without treatment with AM9053. This

sequence contained the catalytic cysteine and has a mass of 1079.5 Da. The spectrum of the

tryptic digest of NAAA after treatment with AM9053 (B, Figure 5.4) exhibited no peak for the

expected combined mass of NAAA and AM9053 (1354.5 Da), while the control peak of 1079.5

Da representing NAAA was undiminished. This indicated there was no covalent activity

associated with the interaction between the peptide containing Cys126 of NAAA and AM9053.

AM9053 was determined to be a competitive inhibitor that binds reversibly within the

active site of NAAA. This was determined by incubation of our fluorogenic substrate PAMCA

with NAAA at various concentrations in the presence and absence of the inhibitor AM9053. This

data was analyzed using a Lineweaver-Burk plot which indicated the competitive nature of

AM9053 (Figure 5.5).

201

Inte

nsity

M/z

Figure 5.4 Tryptic digest of purified human NAAA obtained by MALDI-TOF MS for protein neat (A) and AM9053 treated enzyme (B). The tryptic peptide containing the catalytic nucleophile cysteine is noted with an asterisk (sequence: CTSIVAQDSR, mass: 1079.5 Da). Note that the intensity of the 1079.5 Da peak is undiminished after treatment with AM9053 and the absence of a peak for the covalently modified peptide (expected mass of approximately 1354.5 Da).

*

A

B

*

202

Figure 5.5 Lineweaver-Burk plot analysis of AM9053 inhibition of hNAAA. hNAAA was incubated with the fluorogenic substrate PAMCA at different concentrations in the presence (open cirles) and absence of AM9053 (closed circles) at a concentration of 50 nM. The intersection of the lines at x=0 is indicative of a competitive inhibitor.

5.4 Conclusions

NAEs are known to induce anti-nociceptive and anti-inflammatory responses through

their activation of the PPAR-α receptor. Inhibition of NAAA increases levels of PEA,

oleoylethanolamine, and possibly anandamide, resulting in increased therapeutic benefits. With

a limited availability of NAAA inhibitors we prepared and evaluated various classes of

compounds.

Dual inhibition of endocannabinoid enzymes (FAAH and MGL) has shown promising

results related to drug abuse.37 While inhibition of FAAH and NAAA may have some future

value with overlapping benefits, it is important to identify potent and selective NAAA inhibitors.

With the investigation of NAAA being relatively imprecise, developing ligands as selective

203

inhibitors (as opposed to known ligands that inhibit other cannabinoid metabolizing enzymes) is

important to understand the specific actions of NAAA and its potential as a therapeutic target.

The first attempt at selective NAAA inhibitors used retroamides as it was shown that the

analogs of anandamide containing a retro-amide were metabolically stable in the presence of

FAAH. The synthesized retro-amides exhibited no inhibition of FAAH or MGL, and they were

inactive towards NAAA as well.

The next two classes that we explored were carbonates and carbamates. Utilizing strong

leaving groups, the catalytic cysteine of NAAA could then attack the δ+ carbonyl carbon of the

carbonate or carbamate, resulting in covalent modification of the enzyme which leads to

inhibition of NAAA’s metabolizing actions. The best carbonate derivative (27) had an IC50 of

5.8 μM while retaining selectivity. However, the carbamates were much more potent inhibitors.

The chemical moiety para to the carbamate had a significant effect on NAAA inhibition.

Ligands containing the stronger electron withdrawing group- nitro were better inhibitors

compared to the p-cyano compounds. In addition, the p-nitro compounds were more selective to

NAAA over FAAH. The best compound (38) exhibited an IC50 of 32 nM and 22x selectivity

over FAAH, however, p-nitrophenols as leaving groups can generally be toxic in biological

systems.

We decided to pursue isothiocyanate compounds as covalent inhibitors, as we observed

covalent cysteine-isothiocyanate interactions with the cannabinoid receptors. We identified

pentadecylisothiocyanate (36, IC50 350nM) as being NAAA selective. We continued to reduce

the alkyl chain length and ultimately added a phenyl ring to the tail. This progress gave our best

inhibitor, AM9053 (83) with an IC50 of 39 nM and no activity observed at either FAAH or MGL.

However, our hypothesis that these would behave as covalent modifiers appeared to be incorrect.

204

Fluorescence-based assays did not show a change in slope (fluorescence vs. time) when AM9053

was preincubated with NAAA for either 15 minutes or 3 hours. A reversible mode of action was

also observed when a decrease in inhibition was observed after a rapid dilution assay. This was

supported by a tryptic digest of purified human NAAA- neat and treated with AM9053. No

observable difference in mass spectra of untreated and treated NAAA digests was observed, and

no peak of 1354.5 Da, which would be expected if AM9053 was covalently bound to the

catalytic cysteine nucleophile of NAAA, was present.

This information will focus our continued research on the inhibition of NAAA to pursue

isothiocyanate analogs. Current reported inhibitors of NAAA currently include retroamides

exhibiting micromolar inhibition, and potent, covalent, β-lactone inhibitors that are unstable in

plasma. Multiple tests indicate that these isothiocyanate compounds do not behave through a

covalent mechanism and are selective for NAAA over FAAH and MGL. Isothiocyanates are

stable chemical moieties and are currently very potent NAAA inhibitors. SAR studies of the best

ligands reported here will determine functionalities tolerated by NAAA and possibly improve on

the potency of these inhibitors.

205

5.5 Experimental

NH

O

N-hexadecylcinnamamide, 12, AM9024. Cinnamoyl chloride (22 mg, 0.13 mmol) was added to

a solution of hexadecylamine (32 mg, 0.13 mmol) and catalytic DMAP (3 mg) in CH2Cl2 (4

mL). Upon completion (TLC monitoring) the reaction was washed with water and the organic

layer was separated and filtered through an Isolute® PEAX and SCX-2 column. The filtrate was

collected and the solvent was removed under reduced pressure to yield 12, (15 mg, 32%) as a

white solid. mp 72-75 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.63 (d, J = 15.6 Hz, 1H),

7.50 (dd, J = 1.71, 7.6 Hz, 2H), 7.35 - 7.38 (m, 3H), 6.37 (d, J = 15.6 Hz, 1H), 3.39 (s, 2H), 1.52

- 1.61 (m, 3H), 1.16 - 1.42 (m, 26H), 0.88 (t, J = 6.8 Hz, 3H).

NH

O

O

N-hexadecyl-4-methoxybenzamide, 13, AM9026. Synthesized following the procedure for 12,

(31mg, 33%) white solid. mp 87-88 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.72 (d, J =

8.8 Hz, 2H), 6.92 (d, J = 8.8 Hz, 2H), 5.95 - 6.02 (br. s., 1H), 3.85 (s, 3H), 3.40 - 3.46 (m, 2H),

1.60 (quin, J = 7.3 Hz, 2H), 1.22 - 1.41 (m, 26H), 0.88 (t, J = 6.4 Hz, 2H).

NH

O

N-hexadecyl-2-adamantamide, 14, AM9027. Synthesized following the procedure for 12, (78

mg, 78%) white solid. MP = 78-81 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 5.51 - 5.60

206

(br. s., 1H), 3.19 - 3.26 (m, 2H), 2.04 (m, 3H), 1.84 (d, J = 2.4 Hz, 6H), 1.65 - 1.78 (m, 6H), 1.48

(quin, J = 6.8 Hz, 2H), 1.20 - 1.34 (m, 26H), 0.88 (t, J = 6.8 Hz, 3H).

NH

O

N-hexadecyl-4-methylbenzamide, 15, AM9028. Synthesized following the procedure for 12,

(59 mg, 67%) white solid. mp 75-76 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.65 (d, J =

8.3 Hz, 2H), 7.23 (d, J = 8.3 Hz, 2H), 5.95 - 6.08 (br. s., 1H), 3.38 - 3.48 (m, 2H), 2.39 (s, 3H),


NH

O

F

3-fluoro-N-hexadecylbenzamide, 16, AM9030. Synthesized following the procedure for 12,

(78 mg, 87%) white solid. mp 83-84 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.45 - 7.53

(m, 2H), 7.37 - 7.44 (m, 1H), 7.16 - 7.22 (m, 1H), 5.95 - 6.11 (br. s., 1H), 3.40 - 3.49 (m, 2H),


O

HN

N-hexadecyl-2-(naphthalen-1-yl)acetamide, 17, AM9031. EDCI (148 mg, 0.77 mmol), DMAP

(15 mg, 0.12 mmol), and 1-naphthaleneacetic acid (46 μL, 0.25 mmol) were added to a solution

of hexadecylamine (60 mg, 0.25 mmol) in CH2Cl2 (5 mL) at 0 °C. The reaction stirred under

argon for 6 h. Upon completion the reaction was washed with water and brine. The organic

layer was separated, dried over MgSO4, and removed under reduced pressure. The resulting

residue was chromatographed on silica gel to yield 17 (31 mg, 31%) as a white solid. mp 96-97

207

°C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.94 - 7.98 (m, 1H), 7.87 - 7.91 (m, 1H), 7.84 (d,

J = 8.3 Hz, 1H), 7.54 (ddd, J = 1.7, 5.7, 7.7 Hz, 2H), 7.44 - 7.48 (m, 1H), 7.39 - 7.42 (m, 1H),

5.23 (br. s., 1H), 4.03 (s, 2H), 3.07 - 3.15 (m, 2H), 1.20 - 1.33 (m, 28H), 0.88 (t, J = 7.01 Hz,

3H).

O

NH

O2N

N-hexadecyl-4-nitrobenzamide, 18, AM9033. Synthesized following the procedure for 12,

(300 mg, 82%) white solid. mp 100-101 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 8.29 (d,

J = 8.8 Hz, 2H), 7.92 (d, J = 8.8 Hz, 2H), 6.07 - 6.17 (br. s., 1H), 3.42 - 3.52 (m, 2H), 1.64 (quin,

J = 7.3 Hz, 2H), 1.21 - 1.43 (m, 26H), 0.88 (t, J = 6.8 Hz, 3H).

O

NH

H2N

4-amino-N-hexadecylbenzamide, 19, AM9034. 10% Pd/C (82 mg, 0.077 mmol) was added to a

suspension of 18 (250 mg, 0.64 mmol) in ethyl acetate (20 mL). The reaction was immediately

put under vacuum and flushed with hydrogen (3x). Upon completion the reaction mixture was

filtered and the solvent was removed under reduced pressure. The resulting residue was

chromatographed on silica gel to yield 19 (180 mg, 78%) as a white solid. mp 124-126 °C. 1H

NMR (500 MHz, CHLOROFORM-d) δ 7.59 (d, J = 8.3 Hz, 2H), 6.66 (d, J = 7.8 Hz, 2H), 5.85 -

5.99 (br. s., 1H), 3.93 (br. s., 2H), 3.38 - 3.45 (m, 2H), 1.56 - 1.63 (m, 2H), 1.20 - 1.43 (m, 26H),

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

O

NH

SCN

208

N-hexadecyl-4-isothiocyanatobenzamide, 20, AM9035. 1,1`-thiocarboncyldipyridin-2(1H)-one

(64 μL, 0.84 mmol) was added to a suspension of 19 (100 mg, 0.28 mmol) in anhydrous CH2Cl2

(10 mL) under an atmosphere of argon and stirred for 0.5h. Upon completion the reaction was

washed with water and the organic layer was separated and removed under reduced pressure.

The resulting residue was chromatographed on silica to yield 20 (96 mg, 85%) as a white solid.

mp 85-86 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.75 (d, J = 9.3 Hz, 2H), 7.27 (d, J =

8.8 Hz, 5H), 5.99 - 6.06 (br. s., 1H), 3.41 - 3.47 (m, 2H), 1.61 (quin, J = 7.3 Hz, 2H), 1.19 - 1.41

(m, 26H), 0.87 (t, J = 6.8 Hz, 3H).

O O

ONO2

hexadecyl 4-nitrophenyl carbonate, 25, AM9048. Triethylamine (143 μL, 1.03 mmol), DMAP

(126 mg, 1.03 mmol), and 4-nitrophenyl chloroformate (207 mg, 1.03 mmol) were added to a

solution of 1-hexadecanol (250 mg, 1.03 mmol) in THF (10 mL) stirred at rt. After 2 h the

reaction mixture was diluted with ether and washed with water, brine, and dried over MgSO4.

The solvent was evaporated off under reduced pressure and the resulting residue was

chromatographed on silica to yield 25 (410 mg, 98%) as a white solid. mp 51-52 °C. 1H NMR

(500 MHz, CHLOROFORM-d) δ 8.28 (d, J = 4.88 Hz, 2H), 7.39 (d, J = 4.88 Hz, 2H), 4.29 (t, J

= 6.59 Hz, 2H), 1.72 - 1.80 (m, 2H), 1.22 - 1.47 (m, 26H), 0.88 (t, J = 7.08 Hz, 3H).

O O

ONO2

4-nitrophenyl 8-phenyloctyl carbonate, 26, AM9057. Synthesized following the procedure for

25, (80 mg, 74%) white solid. mp 44-46 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 8.28 (d,

J = 4.9 Hz, 0H), 7.38 (d, J = 4.9 Hz, 0H), 7.27 - 7.30 (m, 2H), 7.14 - 7.21 (m, 3H), 4.28 (t, J =

209

6.8 Hz, 2H), 2.61 (t, J = 7.3 Hz, 2H), 1.75 (quin, J = 6.8 Hz, 4H), 1.58 - 1.66 (m, 2H), 1.38 - 1.45

(m, 2H), 1.35 (m, 6H).

O O

ONO2

4-nitrophenyl 10-phenyldecyl carbonate, 27, AM9056. Synthesized following the procedure

for 25, (72 mg, 71%) white solid. mp 52-54 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 8.28

(d, J = 8.8 Hz, 2H), 7.38 (d, J = 9.3 Hz, 2H), 7.26 - 7.30 (m, 2H), 7.15 - 7.20 (m, 3H), 4.29 (t, J =

6.8 Hz, 2H), 2.60 (t, J = 6.8 Hz, 2H), 1.75 (quin, J = 7.1 Hz, 2H), 1.61 (quin, J = 7.3 Hz, 2H),

1.38 - 1.45 (m, 2H), 1.30 (m, 10H).

NH

O

OCN

4-cyanophenyl dodecylcarbamate, 33, AM9052. 20% Phosgene in toluene (170 μL, 0.34

mmol) was added to a suspension of triethylamine (47 μL, 0.34 mmol) and 4-

hydroxybenzonitrile (40 mg, 0.34 mmol) in anhydrous CH2Cl2 (10 mL) under an atmosphere of

argon at 0 °C. After 45m of stirring, the reaction was treated with dodecylamine (63 mg, 0.34

mmol) and allowed to warm to rt where the reaction stirred for an additional hour. The reaction

was washed with water, brine, and dried over MgSO4. The solvent was evaporated off and the

resulting residue was chromatographed on silica gel to yield 33 (88 mg, 79%) as a white solid.

mp 97-98 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.65 (d, J = 7.8 Hz, 2H), 7.27 (d, J =

7.8 Hz, 2H), 5.06 (br. s., 1H), 3.22 - 3.30 (m, 2H), 1.59 (quin, J = 6.3 Hz, 2H), 1.21 - 1.40 (m,

18H), 0.88 (t, J = 7.1 Hz, 3H).

NH

O

OCN

210

4-cyanophenyl tetradecylcarbamate, 34, AM9051. Synthesized following the procedure for

33, (15 mg, 12%) white solid. mp 102-103 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.65

(d, J = 8.8 Hz, 2H), 7.26 (d, J = 8.8 Hz, 2H), 4.99 - 5.10 (br. s., 1H), 3.24 - 3.30 (m, 2H), 1.59

(quin, J = 6.8 Hz, 2H), 1.26 (s, 22H), 0.88 (t, J = 6.8 Hz, 3H).

NH

O

OCN

4-cyanophenyl pentadecylcarbamate, 35, AM9050. Synthesized following the procedure for


J = 8.8 Hz, 2H), 7.26 (d, J = 8.8 Hz, 2H), 4.98 - 5.11 (m, 1H), 3.23 - 3.30 (m, 2H), 1.57 (quin, J

= 7.1 Hz, 2H), 1.19 - 1.41 (m, 24H), 0.88 (t, J = 6.8 Hz, 3H).

NH

O

OCN

4-cyanophenyl hexadecylcarbamate, 36, AM9049. Synthesized following the procedure for


J = 8.8 Hz, 2H), 7.27 (d, J = 8.8 Hz, 2H), 5.00 - 5.09 (br. s., 1H), 3.25 (s, 2H), 1.57 (quin, J = 7.3

Hz, 2H), 1.20 - 1.41 (m, 26H), 0.88 (t, J = 6.8 Hz, 3H).

N3

(10-azidodecyl)benzene, 37. DBU (144 μL, 0.96 mmol) and DPPA (207 μL, 0.96 mmol) were

added to a solution of 10-phenyl-1-decanol (22, 150 mg, 0.64 mmol) in anhydrous DMF (4 mL)

at 120 °C. The reaction was allowed to stir for 4 h, where it was diluted with ether (10 mL) and

washed with water, brine, and dried over MgSO4. The ether was evaporated off under reduced

pressure and the resulting residue was chromatographed on silica gel to yield 37 (157 mg, 95%)

211

as a pale yellow oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.26 - 7.30 (m, 2H), 7.12 - 7.20

(m, 3H), 3.25 (t, J = 6.8 Hz, 2H), 2.60 (t, J = 6.3 Hz, 2H), 1.55 - 1.65 (m, 4H), 1.21 - 1.40 (m,

12H).

NH

O

ONO2

4-nitrophenyl 10-phenyldecylcarbamate, 38, AM9058. 10% Pd/C (64 mg, 0.06 mmol) was

added to a suspension of 37 (156 mg, 0.60 mmol) in ethyl acetate (10 mL) at rt. The reaction

was placed under vacuum and flushed with hydrogen (3x). Upon completion the catalyst was

filtered off and the remaining solvent removed under reduced pressure. The resulting crude oil

was then diluted in CH2Cl2 (5 mL) and treated with 24 (121 mg, 0.60 mmol) and stirred for 30

m. The reaction mixture was washed with water, brine, and dried over MgSO4. The solvent was

removed and the crude oil was chromatographed on silica gel to yield 38 (32 mg, 13%) as a

white solid. mp 78-80 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 8.24 (d, J = 9.3 Hz, 2H),

7.32 (d, J = 9.3 Hz, 2H), 7.26 - 7.30 (m, 2H), 7.13 - 7.20 (m, 3H), 5.02 - 5.15 (br. s., 1H), 3.23 -

3.33 (m, 2H), 2.60 (t, J = 7.3 Hz, 2H), 1.56 - 1.66 (m, 4H), 1.24 – 1.35 (m, 12H).

NH

O

ONO2

4-nitrophenyl dodecylcarbamate, 40, AM9054. Triethylamine (53 μL, 0.38 mmol), DMAP (12

mg, 0.038 mmol), and 24 (77 mg, 0.38 mmol) were added to a solution of 28 (70 mg, 0.38

mmol) in anhydrous THF (10 mL) under an atmosphere of argon at rt. Upon completion the

reaction was diluted with ether (15 mL), washed with water, brine, and dried over MgSO4. The

solvent was removed under reduced pressure and the resulting residue was chromatographed on

silica gel to yield 40 (131 mg, 98%) as a white solid. mp 96-97 °C. 1H NMR (500 MHz,

212

CHLOROFORM-d) δ 8.25 (d, J = 9.3 Hz, 2H), 7.32 (d, J = 9.3 Hz, 2H), 5.09 (br. s., 1H), 3.24 -

3.32 (m, 2H), 1.59 (quin, J = 7.1 Hz, 2H), 1.22 - 1.40 (m, 18H), 0.88 (t, J = 6.6 Hz, 3H).

NH

O

ONO2O

4-nitrophenyl 4-(benzyloxy)phenylcarbamate, 41, AM9072. Synthesized following the

procedure for 40, (37 mg, 32%) white solid. MP = 156-159 °C. 1H NMR (500 MHz,

CHLOROFORM-d) δ 8.28 (d, J = 8.8 Hz, 2H), 7.29 - 7.46 (m, 9H), 6.98 (d, J = 9.3 Hz, 2H),

6.89 - 6.94 (m, 1H), 5.06 (s, 2H).

NH

O

O

Naphthalen-2-yl 10-phenyldecylcarbamate, 49, AM9061. DMAP (5mg, 0.04 mmol) and 22

(15 mg, 0.64 mmol) were added to a solution of 2-naphthyl chloroformate (21 mg, 0.1 mmol) in

CH2Cl2 (3 mL) at 0 °C. The reaction was allowed to warm to room temperature and stir for 16h.

Upon completion the reaction mixture was filtered through an Isolute® SCX-2 column. The

collected solvent was removed under reduced pressure. The resulting residue was

chromatographed on silica gel to yield 49 (26mg, 100%) as a colorless oil. 1H NMR (500 MHz,

CHLOROFORM-d) δ 7.91 - 7.98 (m, 1H), 7.82 - 7.88 (m, 1H), 7.71 (d, J = 8.3 Hz, 1H), 7.38 -

7.52 (m, 4H), 7.25 - 7.33 (m, 2H), 7.18 (m, Hz, 3H), 5.22 (br. s., 1H), 3.24 - 3.38 (m, 2H), 2.60

(t, J = 7.8 Hz, 2H), 1.61 (quin, J = 7.3 Hz, 4H), 1.27 - 1.41 (m, 12H).

NH

O

OF

213

4-fluorophenyl 10-phenyldecylcarbamate, 50, AM9062. Synthesized following the procedure

for 49, (24mg, 100%) white solid. mp = 65-66 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ

7.24 - 7.30 (m, 3H), 7.16 - 7.19 (m, 2H), 7.05 - 7.10 (m, 2H), 6.99 - 7.05 (m, 2H), 4.94 - 5.02 (br.

s., 1H), 3.22 - 3.28 (m, 2H), 2.60 (t, J = 6.8 Hz, 2H), 1.53 - 1.65 (m, 4H), 1.25 - 1.35 (m, 12H).

NH

O

OCl

2-chlorophenyl 10-phenyldecylcarbamate, 51, AM9063. Synthesized following the procedure

for 49, (14 mg, 56%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.39 - 7.47 (m,

2H), 7.22 - 7.30 (m, 3H), 7.12 - 7.21 (m, 4H), 5.11 (br. s., 1H), 3.22 - 3.32 (m, 2H), 2.60 (t, J =

7.3 Hz, 2H), 1.56 - 1.66 (m, 4H), 1.19 - 1.44 (m, 12H).

NH

O

O

CF3

3-(trifluoromethyl)phenyl 10-phenyldecylcarbamate, 52, AM9064. Synthesized following the

procedure for 49, (10 mg, 37%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.43 -

7.49 (m, 2H), 7.41 (s, 1H), 7.30 - 7.35 (m, 1H), 7.26 - 7.30 (m, 2H), 7.15 - 7.19 (m, 3H), 5.05

(br. s., 1H), 3.22 - 3.30 (m, 2H), 2.60 (t, J = 7.3 Hz, 2H), 1.56 - 1.66 (m, 4H), 1.22 - 1.41 (m,

12H).

NH

O

O

NO2

2-nitrophenyl 10-phenyldecylcarbamate, 53, AM9065. Synthesized following the procedure

for 49, (12 mg, 46%) white solid. mp = 33-36 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ

8.04 (dd, J = 1.5, 8.3 Hz, 1H), 7.62 (dt, J = 1.5, 7.8 Hz, 1H), 7.33 - 7.39 (m, 2H), 7.24 - 7.32 (m,

214

3H), 7.12 - 7.21 (m, 2H), 5.19 (br. s., 1H), 3.22 - 3.31 (m, 2H), 2.60 (t, J = 7.8 Hz, 2H), 1.51 -

1.66 (m, 4H), 1.21 - 1.43 (m, 12H).

NH

O

OOMe

4-methoxyphenyl 10-phenyldecylcarbamate, 54, AM9066. Synthesized following the

procedure for 49, (13 mg, 52%) white solid. mp = 68-69 °C. 1H NMR (500 MHz,

CHLOROFORM-d) δ 7.23 - 7.30 (m, 2H), 7.14 - 7.20 (m, 3H), 7.00 - 7.06 (m, 2H), 6.86 (d, J =

9.3 Hz, 2H), 4.95 (br. s., 1H), 3.79 (s, 3H), 3.18 - 3.29 (m, 2H), 2.60 (t, J = 7.3 Hz, 2H), 1.50 -

1.64 (m, 4H), 1.22 - 1.38 (m, 12H).

NH

O

OBr

4-bromophenyl 10-phenyldecylcarbamate, 55, AM9067. Synthesized following the procedure

for 49, (7 mg, 25%) white solid. mp = 73-74 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.41

- 7.48 (m, 2H), 7.26 - 7.30 (m, 2H), 7.12 - 7.21 (m, 3H), 6.97 - 7.05 (m, 2H), 4.99 (br. s., 1H),

3.21 - 3.28 (m, 2H), 2.60 (t, J = 7.8 Hz, 2H), 1.48 - 1.66 (m, 4H), 1.22 - 1.40 (m, 12H).

O

NH

OH

Palmitoylethanolamide, 57: EDCI (673 mg, 3.51 mmol), DMAP (44 mg, 0.35 mmol), and

ethanolamine (0.141 mL, 2.34 mmol) were added to a solution palmitic acid 56 (300 mg, 1.17

mmol) was stirred in 10 mL of anhydrous CH2Cl2 at 0 oC. The reaction was allowed to stir under

argon for 4 hours while warming to room temperature. Upon completion the reaction mixture

was diluted with CH2Cl2, washed with water and brine. The organic layer was collected and

concentrated. The resulting residue was chromatographed on silica to yield 57 (312 mg, 72%) as

215

a white solid. mp is in agreement with literature values.37 1H NMR (500 MHz,

CHLOROFORM-d) δ 3.70 - 3.76 (m, 2H), 3.43 (q, J = 5.3 Hz, 2H), 2.17 - 2.24 (m, 2H), 1.64

(quin, J = 7.5 Hz, 2H), 1.52 (d, J = 1.0 Hz, 1H), 1.19 - 1.35 (m, 26H).

O

NH

N3

N-(2-azidoethyl)palmitamide, 58: Synthesized following the procedure for 37, (36 mg, 66%)

off-white solid. mp 68-69 °C. 1H NMR (500 MHz, CHLOROFORM-d) δ 3.35 - 3.49 (m, 4H),

2.19 (t, J = 7.3 Hz, 2H), 1.63 (quin, J = 7.5 Hz, 2H), 1.19 - 1.37 (m, 24H), 0.88 (t, J = 6.8 Hz,

3H).

O

NH

NCS

N-(2-isothiocyanatoethyl)palmitamide, 59, AM9019: PPh3 (36 mg, 0.14 mmol) and CS2 (10

μL, 0.16 mmol) were added to a solution of 58 (35 mg, 0.11 mmol) in anhydrous THF (5 mL)

under an atmosphere of argon. The reaction was allowed to stir for 48 h. The reaction mixture

was concentrated and the resulting residue was chromatographed on silica gel to yield 59 (25 mg,

68%) as a white solid. mp 72-74 °C.1H NMR (500 MHz, CHLOROFORM-d) δ 5.79 - 5.99 (m,

1H), 3.68 (t, J = 5.4 Hz, 2H), 3.51 (q, J = 5.9 Hz, 2H), 2.22 (t, J = 7.6 Hz, 2H), 1.64 (td, J = 7.5,

14.4 Hz, 2H), 1.22 - 1.39 (m, 24H), 0.88 (t, J = 7.1 Hz, 3H). IR (neat) cm-1 3288, 2917, 2849,

2182, 2094, 1645. HRMS for C19H36N2OS (MH+) 341.2623. Calcd. 341.2627.

NCS

1-isothiocyanatopentadecane, 66, AM9023: Synthesized following the procedure for 20, (55

mg, 94%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 3.51 (t, J = 6.8 Hz, 2H), 1.69

216

(td, J = 6.8, 15.1 Hz, 2H), 1.36 - 1.44 (m, 2H), 1.21 - 1.34 (m, 22H), 0.88 (t, J = 6.8 Hz, 3H). IR

(neat) cm-1 2924, 2854, 2185, 2090. HRMS for C16H30NS (M-H+) 268.2111. Calcd. 268.2099.

NCS

1-isothiocyanatotetradecane, 67, AM9046: Synthesized following the procedure for 20, (72

mg, 98%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 3.51 (t, J = 6.6 Hz, 2H), 1.69

(td, J = 6.8, 15.1 Hz, 2H), 1.36 - 1.45 (m, 2H), 1.22 - 1.35 (m, 20H), 0.88 (t, J = 6.8 Hz, 3H). IR


NCS

1-isothiocyanatododecane, 68, AM9045: Synthesized following the procedure for 20, (71 mg,

97%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 3.51 (t, J = 6.6 Hz, 2H), 1.69 (td,

J = 6.8, 15.1 Hz, 2H), 1.37 - 1.44 (m, 2H), 1.22 - 1.35 (m, 16H), 0.88 (t, J = 7.1 Hz, 3H). IR


NCS

1-isothiocyanatodecane, 69, AM9042: Synthesized following the procedure for 20, (62 mg,

98%) yellow oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 3.51 (t, J = 6.6 Hz, 2H), 1.69 (tt, J =

6.4, 7.6 Hz, 2H), 1.36 - 1.45 (m, 2H), 1.19 - 1.35 (m, 12H), 0.88 (t, J = 7.0 Hz, 3H). IR (neat)

cm-1 2923, 2854, 2182, 2083. HRMS for C11H20NS (M-H+) 198.1320. Calcd. 198.1316.

NCS

2-isothiocyanatoundecane, 70, AM9043: The procedure for 10a was used to synthesize 10e (56

mg, 90%) as an yellow oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 3.76 (dqd, J = 5.3, 6.6,

8.2 Hz, 1H), 1.59 - 1.68 (m, 1H), 1.51 - 1.59 (m, 1H), 1.35 (d, J = 6.4 Hz, 3H), 1.23 - 1.33 (m,

14H), 0.89 (t, J = 6.7 Hz, 3H). IR (neat) cm-1 2924, 2854, 2184, 2082. HRMS for C12H23NS (M+)

213.1562. Calcd. 213.1551.

217

SO

NCS

DL-sulforane, 71, AM9044: Purchased from Sigma Aldrich.

N3

(4-azidobutyl)benzene, 75: Synthesized following the procedure for 37, (42 mg, 53%) pale

yellow oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.27 - 7.32 (m, 2H), 7.13 - 7.23 (m, 3H),

3.29 (t, J = 6.8 Hz, 2H), 2.65 (t, J = 7.6 Hz, 2H), 1.68 - 1.76 (m, 2H), 1.60 - 1.67 (m, 2H).

N3

(5-azidopentyl)benzene, 76: Synthesized following the procedure for 37, (109 mg, 92%) pale


3.26 (t, J = 6.8 Hz, 2H), 2.62 (t, J = 7.6 Hz, 2H), 1.57 - 1.70 (m, 4H), 1.37 - 1.46 (m, 2H).

N3

(6-azidohexyl)benzene, 77: Synthesized following the procedure for 37, (99 mg, 91%) pale

yellow oil. 1H NMR (500 MHz, CHLOROFORM-d) d 7.26 - 7.31 (m, 2H), 7.14 - 7.22 (m, 3H),

3.25 (t, J = 6.8 Hz, 2H), 2.61 (t, J = 7.6 Hz, 2H), 1.56 - 1.68 (m, 4H), 1.32 - 1.45 (m, 4H).

N3

(8-azidooctyl)benzene, 78: Synthesized following the procedure for 37, (136 mg, 81%) pale


3.25 (t, J = 7.1 Hz, 2H), 2.55 - 2.64 (m, 2H), 1.55 - 1.67 (m, 4H), 1.26 - 1.41 (m, 8H).

NCS

218

(4-isothiocyanatobutyl)benzene, 79, AM9038: Synthesized following the procedure for 59, (41

mg, 90%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.27 - 7.32 (m, 2H), 7.14 -

7.23 (m, 3H), 3.52 (t, J = 6.4 Hz, 2H), 2.66 (t, J = 7.1 Hz, 2H), 1.67 - 1.81 (m, 4H). IR (neat) cm-

1 3026, 2926, 2859, 2183, 2089. HRMS for C11H13NS (M+) 191.0760. Calcd. 191.0769.

NCS

(5-isothiocyanatopentyl)benzene, 80, AM9037: Synthesized following the procedure for 59,

(116 mg, 90%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.25 - 7.29 (m, 2H),

7.15 - 7.22 (m, 3H), 3.50 (t, J = 6.6 Hz, 2H), 2.63 (t, J = 7.3 Hz, 2H), 1.72 (td, J = 6.8, 15.1 Hz,

2H), 1.66 (td, J = 7.6, 15.5 Hz, 2H), 1.42 - 1.50 (m, 2H). IR (neat) cm-1 3026, 2935, 2857, 2183,

2086. HRMS for C12H15NS (M+) 205.0938. Calcd. 205.0925.

NCS

(6-isothiocyanatohexyl)benzene, 81, AM9036: Synthesized following the procedure for 59, (72


7.21 (m, 3H), 3.50 (t, J = 6.6 Hz, 2H), 2.62 (t, J = 7.8 Hz, 2H), 1.60 - 1.73 (m, 4H), 1.40 - 1.48

(m, 2H), 1.32 - 1.40 (m, 2H). IR (neat) cm-1 3026, 2931, 2856, 2183, 2092. HRMS for C13H17NS

(M+) 219.1073. Calcd. 219.1082.

NCS

(8-isothiocyanatooctyl)benzene, 82, AM9047: Synthesized following the procedure for 59, (46


7.21 (m, 3H), 3.50 (t, J = 6.8 Hz, 2H), 2.60 (t, J = 6.8 Hz, 2H), 1.58 - 1.72 (m, 4H), 1.37 - 1.44

219

(m, 2H), 1.28 - 1.37 (m, 6H). IR (neat) cm-1 3026, 2927, 2955, 2179, 2090. HRMS for C15H21NS

(M+) 247.1387. Calcd. 247.1395.

NCS

(10-isothiocyanatodecyl)benzene, 83, AM9053: Synthesized following the procedure for 59,

(74 mg, 73%) colorless oil. 1H NMR (500 MHz, CHLOROFORM-d) δ 7.24 - 7.31 (m, 2H), 7.14

- 7.21 (m, 3H), 3.50 (t, J = 6.6 Hz, 2H), 2.60 (t, J = 7.3 Hz, 2H), 1.65 - 1.72 (m, 2H), 1.61 (quin,

J = 7.5 Hz, 2H), 1.36 - 1.44 (m, 2H), 1.24 - 1.35 (m, 10H). IR (neat) cm-1 3026, 2924, 2854,

2178, 2088. HRMS for C17H25NS (M+) 275.1715. Calcd. 275.1708.

O

O

Methyl 7-(naphthalen-2-yl)hept-6-ynoate, 85: A mixture of 2-bromonaphthalene 84 (400 mg,

1.93 mmol), methyl 6-heptynoate (0.282 mL, 1.93 mmol), triethylamine (0.080 mL, 0.57 mmol),

CuI (43 mg, 0.23 mmol), and bis(triphenylphosphine)palladium(II)dichloride (84 mg, 0.12

mmol) in anhydrous DMF (3 mL) were irradiated in a microwave for 20 min at 100 °C. Upon

completion the reaction was diluted with ether and washed with water, brine and dried over

MgSO4. The organic layer was evaporated off under reduced pressure and the resulting residue

was chromatographed on silica to yield 85, (172 mg, 32%) as a colorless oil. 1H NMR (500

MHz, CHLOROFORM-d) δ 7.90 (s, 1H), 7.72 - 7.83 (m, 3H), 7.42 - 7.51 (m, 3H), 3.69 (s, 3H),

2.49 (t, J = 6.8 Hz, 2H), 2.40 (t, J = 7.3 Hz, 2H), 1.85 (quin, J = 7.8 Hz, 2H), 1.69 (quin, J = 7.3

Hz, 2H).

220

O

O

Methyl 7-(naphthalene-2-yl)heptanoate, 86: Pd/C (16 mg, 0.015 mmol) was added to a

solution of 85 (172 mg, 0.60 mmol) in ethyl acetate (10 mL). The solution was placed under

vacuum and flushed with H2 (3x). Upon completion (monitored by TLC) the reaction was

filtered through celite, and the solvent was evaporated off under reduced pressure. The resulting

residue was chromatographed on silica gel to yield 86 (168 mg, 98%) as a colorless oil. 1H NMR

(400 MHz, CHLOROFORM-d) δ 7.73 - 7.83 (m, 3H), 7.60 (s, 1H), 7.37 - 7.48 (m, 2H), 7.32

(dd, J = 1.8, 8.4 Hz, 1H), 3.66 (s, 3H), 2.76 (t, J = 7.3 Hz, 2H), 2.30 (t, J = 7.7 Hz, 2H), 1.67 -

1.75 (m, 2H), 1.58 - 1.66 (m, 2H), 1.31 - 1.43 (m, 4H).

OH

7-(naphthalene-2-yl)heptan-1-ol, 87: A 1.0M solution of LiAlH4 in THF (1.24 mL, 1.24 mmol)

was added a solution of 86 (168 mg, 0.62 mmol) in anhydrous THF (5 mL) at 0 oC. The reaction

mixture was stirred under argon and slowly warmed to room temperature where it stirred for an

additional hour. Once the reaction was complete a 1.0M HCl solution was added drop wise until

the reaction mixture was slightly acidic. The lipophilic products were extracted with Et2O, and

the aqueous layer was washed with Et2O three additional times. The combined layers were dried

with MgSO4 and concentrated to give 87 (130 mg, 87%) as a yellowish wax. 1H NMR (400

MHz, CHLOROFORM-d) δ 7.73 - 7.83 (m, 3H), 7.61 (s, 1H), 7.37 - 7.48 (m, 2H), 7.29 - 7.35

(m, 1H), 3.59 - 3.68 (m, 2H), 2.77 (t, J = 7.3 Hz, 2H), 1.66 - 1.76 (m, 2H), 1.52 - 1.61 (m, 2H),

1.31 - 1.42 (m, 6H).

N3

221

2-(7-azidoheptyl)naphthalene, 88: Synthesized following the procedure for 37, (118 mg, 82%)

pale yellow oil. 1H NMR (400 MHz, CHLOROFORM-d) δ 7.73 - 7.84 (m, 3H), 7.61 (s, 1H),

7.38 - 7.49 (m, 2H), 7.33 (d, J = 8.1 Hz, 1H), 3.25 (t, J = 7.0 Hz, 2H), 2.77 (t, J = 7.7 Hz, 2H),

1.66 - 1.77 (m, 2H), 1.57 - 1.64 (m, 2H), 1.37 (br. s., 6H).

NCS

2-(7isothiocyanatoheptyl)naphthalene, 89, AM9060: Synthesized following the procedure for

59, (41 mg, 77%) yellow oil.1H NMR (500 MHz, CHLOROFORM-d) δ 7.72 - 7.82 (m, 4H),

7.60 (s, 1H), 7.37 - 7.48 (m, 3H), 7.32 (dd, J = 1.5, 8.3 Hz, 1H), 3.47 (t, J = 6.8 Hz, 2H), 2.77 (t,

J = 7.6 Hz, 2H), 1.62 - 1.75 (m, 4H), 1.31 - 1.43 (m, 6H). IR (neat) cm-1 3053, 2929, 2856, 2093.

HRMS for C18H21NS (MH+) 284.1478. Calcd. 284.1473.

O

NH

OH

N-(2-hydroxyethyl)-7-(naphthalen-2-yl)heptanamide, 91: EDCI (357 mg, 1.86 mmol), DMAP

(15 mg, 0.12 mmol), and ethanolamine (53 μL, 0.87 mmol) were added to a solution 7-

(naphthalen-2-yl)heptanoic acid 90 (159 mg, 0.62 mmol) stirred in anhydrous CH2Cl2 (10 mL)

at 0 oC. The reaction was allowed to stir under argon for 4 hours while warming to room

temperature. Upon completion the reaction mixture was diluted with CH2Cl2, washed with water

and brine. The organic layer was collected and concentrated. The resulting oil was

chromatographed on silica gel to yield 91 (93 mg, 50%) as a white solid. 1H NMR (500 MHz,

CHLOROFORM-d) δ 7.73 - 7.82 (m, 3H), 7.60 (s, 1H), 7.38 - 7.48 (m, 2H), 7.32 (dd, J = 1.5,

8.3 Hz, 1H), 5.88 (br. s., 1H), 3.70 (t, J = 5.4 Hz, 2H), 3.40 (q, J = 5.4 Hz, 2H), 2.77 (t, J = 7.8

Hz, 2H), 2.15 - 2.21 (m, 2H), 1.68 - 1.75 (m, 2H), 1.60 - 1.68 (m, 1H), 1.32 - 1.43 (m, 4H).

222

O

HN

NCS

N-(2-isothiocyanatoethyl)-7-(naphthalene-2-yl)heptanamide, 93, AM9071: Synthesized

following the procedures for 58 and 59, (22 mg, 34%) white solid. mp 67-70 °C. 1H NMR (500

MHz, CHLOROFORM-d) δ 7.89 - 7.92 (m, 1H), 7.71 - 7.82 (m, 4H), 7.41 - 7.50 (m, 2H), 3.68

(t, J = 5.4 Hz, 2H), 3.52 (q, J = 5.9 Hz, 2H), 2.51 (t, J = 6.8 Hz, 2H), 2.32 (t, J = 7.8 Hz, 2H),

1.88 (quin, J = 7.8 Hz, 3H), 1.67 - 1.77 (m, 4H), 1.48 - 1.55 (m, 2H). IR (neat) cm-1 3303, 3058,

2930, 2856, 2198, 2110, 1652. HRMS for C20H24N2OS (M+) 340.1584. Calcd. 340.1609.

NNCS

4-(4-isothiocyanatobenzyl)pyridine, 102, AM10318: Synthesized following the procedure for

20, (31 mg, 86%) orange solid. MP = 53-54 °C. 1H NMR (399 MHz, CHLOROFORM-d) δ 8.50

- 8.53 (m, 2H), 7.13 - 7.20 (m, 4H), 7.05 - 7.09 (m, 2H), 3.96 (s, 2H). IR (neat) cm-1 3067, 3027,

2932, 2174, 2088, 1597. HRMS for C13H10N2S (MH+) 227.0637. Calcd. 227.0643.

NCSN

N

1-isothiocyanato-2-(4-benzylpiperazino)ethane, 103, AM10323: Synthesized following the

procedure for 20, (32 mg, 90%) yellow oil. 1H NMR (399 MHz, CHLOROFORM-d) δ 7.28 -

7.37 (m, 4H), 7.21 - 7.28 (m, 1H), 3.58 (t, J = 6.2 Hz, 2H), 3.51 (s, 2H), 2.67 (t, J = 6.6 Hz, 2H),

2.45 - 2.57 (m, 8H). IR (neat) cm-1 3027, 2939, 2810, 2190, 2097. HRMS for C14H19N3S (MH+)

262.1371. Calcd. 262.1378.

NCS

ON

223

4-(4-(isothiocyanatomethyl)phenyl)morpholine, 104, AM10324: Synthesized following the

procedure for 20, (5.4 mg, 90%) colorless oil. 1H NMR (399 MHz, CHLOROFORM-d) δ 7.19 -

7.28 (m, 2H), 6.88 - 6.96 (m, 2H), 4.62 (s, 2H), 3.83 - 3.90 (m, 4H), 3.13 - 3.24 (m, 4H). IR

(neat) cm-1 3031, 2957, 2856, 2172, 2090, 1724. HRMS for C12H14N2OS (MH+) 235.0902.

Calcd. 235.0905.

NCSN

4-benzyl-1-(2-isothiocyanatoethyl)piperidine, 105, AM10325: Synthesized following the

procedure for 20, (5.2 mg, 87%) colorless oil. 1H NMR (399 MHz, CHLOROFORM-d) δ 7.24 -

7.32 (m, 2H), 7.16 - 7.22 (m, 1H), 7.14 (d, J = 7.3 Hz, 2H), 3.57 (t, J = 6.2 Hz, 2H), 2.85 (d, J =

11.7 Hz, 2H), 2.64 (t, J = 6.6 Hz, 2H), 2.53 (d, J = 6.6 Hz, 2H), 2.03 (dt, J = 2.6, 11.5 Hz, 2H),

1.59 - 1.71 (m, 2H), 1.47 - 1.56 (m, 1H), 1.23 - 1.37 (m, 2H). IR (neat) cm-1 3026, 2928, 2804,

2190, 2101. HRMS for C15H20N2S (MH+) 261.1414. Calcd. 261.1425.

NCSON

4-(morpholinomethyl)benzylisothiocyanate, 106, AM10326: Synthesized following the

procedure for 20, (5.7 mg, 95%) colorless oil. 1H NMR (399 MHz, CHLOROFORM-d) δ 7.37

(s, 2H), 7.23 - 7.29 (m, 2H), 4.70 (s, 2H), 3.67 - 3.76 (m, 4H), 3.50 (s, 2H), 2.45 (d, J = 4.4 Hz,

4H). IR (neat) cm-1 3030, 2961, 2854, 2162, 2084, 1612. HRMS for C13H16N2OS (MH+)

249.1052. Calcd. 249.1062.

N

O

ONCS

Benzyl 4-(isothiocyanatomethyl)tetrahydro-1(2H)-pyridinecarboxylate, 107, AM10327:

Synthesized following the procedure for 20, (5.0 mg, 86%) colorless oil. 1H NMR (500 MHz,

224

CHLOROFORM-d) δ 7.30 - 7.39 (m, 5H), 5.13 (s, 2H), 4.14 - 4.41 (m, 2H), 3.42 (d, J = 6.4 Hz,

2H), 2.79 (br. s., 2H), 1.80 - 1.92 (m, 1H), 1.76 (d, J = 13.2 Hz, 2H), 1.16 - 1.33 (m, 2H). IR

(neat) cm-1 3027, 2934, 2855, 2187, 2097, 1697. HRMS for C15H18N2O2S (MH+) 291.1175.

Calcd. 291.1167.

NCS

ON

N

2-(4-(isothiocyanatomethyl)phenoxy)-6-methylpyrazine, 108, AM10328: Synthesized

following the procedure for 20, (5.3 mg, 90%) colorless oil. 1H NMR (399 MHz,

CHLOROFORM-d) δ 8.18 (s, 2H), 7.36 (d, J = 8.8 Hz, 2H), 7.12 - 7.23 (m, 2H), 4.74 (s, 2H),

2.43 (s, 4H). IR (neat) cm-1 3048, 2926, 2852, 2172, 2090. HRMS for C13H11N3OS (MH+)

258.0697. Calcd. 258.0701.

NCSO

1-(isothiocyanatomethyl)-4-(phenoxymethyl)benzene, 19, AM10329: Synthesized following

the procedure for 20, (5.0 mg, 86%) colorless oil. 1H NMR (399 MHz, CHLOROFORM-d) δ

8.18 (s, 2H), 7.36 (d, J = 8.8 Hz, 2H), 7.12 - 7.23 (m, 2H), 4.74 (s, 2H), 2.43 (s, 4H). IR (neat)

cm-1 3058, 3032, 2925, 2856, 2172, 2091, 1723, 1598. HRMS for C15H13NOS (MH+) 255.0728.

Calcd. 255.0718.

MALDI-TOF MS analysis of hNAAA inhibition by AM9053

Ligand-assisted protein structure (LAPS) type analysis was used to determine if AM9053

covalently modified NAAA.38 This approach consists of incubating the purified enzyme alone

and with AM9053, evaluating extent of inactivation, performing a tryptic digest, comparing the

225

peptide profile fingerprints using MALDI-TOF MS, and then assigning the site and nature of any

covalent modification by MS/MS analysis.

Fluorometric assay to determine hNAAA inhibition using N-(4-methyl

coumarin)palmitamide (PAMCA) substrate. We previously described the fluorogenic

substrate N-(4-methyl coumarin)palmitamide (PAMCA), which is hydrolyzed by NAAA to the

fluorescent compound 7-amino-4-methyl coumarin (AMC) and palmitic acid.39 For hNAAA

inhibition we conducted three point concentration assays with compounds to determine their

potencies and ranges of enzyme inhibition. Purified activated NAAA (final concentration of 0.25

µg/mL) was incubated in assay buffer39 made up to a total volume of 180 µL, followed by

addition of the compound dissolved in 10 µL DMSO (along with DMSO neat for the control

sample) with the final concentrations for each compound of 1, 10, and 100 µM, in triplicate on a

96 well plate. These samples were allowed to incubate for 15 min at room temperature and then

10 µL of a PAMCA stock solution in DMSO (final PAMCA concentration 10 µM) was added.

After 5 minutes of agitation on a shaking plate, the reaction was allowed to proceed at 37 °C for

30 minutes and enzyme activity was monitored and calculated as previously described.39

For compounds that inhibited hNAAA in the range of IC50 < 1 µM full inhibition curves

using eight different concentrations of inhibitor (8 point assay) were generated. To set up 8 point

fluorescent assay for each point, the compound in 45 µL DMSO and purified activated NAAA

(final enzyme concentration of 0.25 µg/mL) in 810 µL of NAAA assay buffer were incubated for

3 hours in order for the covalent compounds to reach full inhibition. For the fluorescent assay,

190 µL of each of the above samples (in triplicate) were transferred to a 96 well plate, followed

by addition of 10 µL of a PAMCA stock solution in DMSO for a final PAMCA concentration of

10 µM. After 5 minutes of agitation on a shaking plate, the reaction was allowed to proceed at

226

37 °C for 30 minutes and enzyme activity was monitored and calculated as previously

described.39

Trypsin digestion of hNAAA treated with inhibitors. To 10 µg (0.2 nmol) of purified and

activated NAAA in 18 µL of 100 mM citrate-sodium phosphate buffer (pH 4.5) 2 µL of a

DMSO solution containing 2 nmol of the compound of interest were added or 2 µL DMSO. The

inhibitor and DMSO treated enzyme solutions were incubated at 37 °C for 2 hours and then

desalted prior to digestion. These were desalted by re-concentrating 3 times to original volume

after 25 fold dilution with 50 mM ammonium bicarbonate buffer, pH 8.0, using 10 kDa

membrane Ultra-0.5 Centrifugal Filters (Millipore). The NAAA samples were incubated

overnight at 37 °C with MS-grade trypsin (“Trypsin Gold”, Promega) at a NAAA:trypsin mass

to mass ratio of 100:1. The tryptic digested NAAA was analyzed immediately or frozen at -80

°C for future analysis.

MALDI-TOF-MS Analysis. 0.5 µL of the trypsin digested NAAA was mixed with 0.5 µL α-

cyano-4-hydroxycinnaminic acid matrix solution (5 mg/mL dissolved in 50% acetonitrile, 50%

water, and 0.1% trifluoroacetic acid) and spotted onto an Opti-TOF 384-well plate insert.

MALDI-TOF MS spectra were acquired on a 4800 MALDI TOF/TOF mass spectrometer

(Applied Biosystems, Foster City, CA) fitted with a 200-Hz solid state UV laser (wavelength 355

nm). The spectra of the peptides were acquired in reflectron mode. The conditions used for the

MS experiments and instrument calibration were performed as described by Zvonok et. al.40

227

5.6 References

1. Coulon, D.; Faure, L.; Salmon, M.; Wattelet, V.; Bessoule, J.-J. N-Acylethanolamines and related compounds: Aspects of metabolism and functions. Plant Science 2012, 184, 129-140. 2. Kobilka, B. K. G protein coupled receptor structure and activation. Biochimica et Biophysica Acta (BBA) - Biomembranes 2007, 1768, 794-807. 3. Smita, K.; Kumar, V. S.; Premendran, J. S. Anandamide: an update. Fundamental & clinical Pharmacology 2007, 1-8. 4. Lo Verme, J.; Fu, J.; Astarita, G.; La Rana, G.; Russo, R.; Calignano, A.; Piomelli, D. The Nuclear Receptor Peroxisome Proliferator-Activated Receptor-α Mediates the Anti-Inflammatory Actions of Palmitoylethanolamide. Molecular Pharmacology 2005, 67, 15-19. 5. Dhake, K. P.; Qureshi, Z. S.; Singhal, R. S.; Bhanage, B. M. Candida antarctica lipase B-catalyzed synthesis of acetamides using [BMIm(PF6)] as a reaction medium. Tetrahedron Letters 2009, 50, 2811-2814. 6. Han, C.; Lee, J. P.; Lobkovsky, E.; Porco, J. A. Catalytic Ester−Amide Exchange Using Group (IV) Metal Alkoxide−Activator Complexes. Journal of the American Chemical Society 2005, 127, 10039-10044. 7. Godlewski, G.; Offertáler, L.; Wagner, J. A.; Kunos, G. Receptors for acylethanolamides—GPR55 and GPR119. Prostaglandins & Other Lipid Mediators 2009, 89, 105-111. 8. LoVerme, J.; La Rana, G.; Russo, R.; Calignano, A.; Piomelli, D. The search for the palmitoylethanolamide receptor. Life Sciences 2005, 77, 1685-98. 9. Lambert, D. M.; Vandevoorde, S.; Jonsson, K.-O.; Fowler, C. J. The Palmitoylethanolamide Family: A New Class of Anti-Inflammatory Agents? Current Medicinal Chemistry 2002, 9, 663-674. 10. Clement, A. B.; Hawkins, E. G.; Lichtman, A. H.; Cravatt, B. F. Increased seizure susceptibility and proconvulsant activity of anandamide in mice lacking fatty acid amide hydrolase. J Neurosci 2003, 23, 3916-23. 11. Cravatt, B. F.; Demarest, K.; Patricelli, M. P.; Bracey, M. H.; Giang, D. K.; Martin, B. R.; Lichtman, A. H. Supersensitivity to anandamide and enhanced endogenous cannabinoid signaling in mice lacking fatty acid amide hydrolase. Proceedings of the National Academy of Sciences of the United States of America 2001, 98, 9371-9376. 12. Lichtman, A. H.; Hawkins, E. G.; Griffin, G.; Cravatt, B. F. Pharmacological activity of fatty acid amides is regulated, but not mediated, by fatty acid amide hydrolase in vivo. J Pharmacol Exp Ther 2002, 302, 73-9. 13. Lichtman, A. H.; Leung, D.; Shelton, C. C.; Saghatelian, A.; Hardouin, C.; Boger, D. L.; Cravatt, B. F. Reversible inhibitors of fatty acid amide hydrolase that promote analgesia: evidence for an unprecedented combination of potency and selectivity. J Pharmacol Exp Ther 2004, 311, 441-8. 14. Fegley, D.; Gaetani, S.; Duranti, A.; Tontini, A.; Mor, M.; Tarzia, G.; Piomelli, D. Characterization of the fatty acid amide hydrolase inhibitor cyclohexyl carbamic acid 3'-carbamoyl-biphenyl-3-yl ester (URB597): effects on anandamide and oleoylethanolamide deactivation. J Pharmacol Exp Ther 2005, 313, 352-8. 15. Sun, Y. X.; Tsuboi, K.; Zhao, L. Y.; Okamoto, Y.; Lambert, D. M.; Ueda, N. Involvement of N-acylethanolamine-hydrolyzing acid amidase in the degradation of anandamide

228

and other N-acylethanolamines in macrophages. Biochimica Et Biophysica Acta 2005, 1736, 211-20. 16. Tsuboi, K.; Sun, Y. X.; Okamoto, Y.; Araki, N.; Tonai, T.; Ueda, N. Molecular characterization of N-acylethanolamine-hydrolyzing acid amidase, a novel member of the choloylglycine hydrolase family with structural and functional similarity to acid ceramidase. Journal of Biological Chemistry 2005, 280, 11082-11092. 17. Ueda, N.; Yamanaka, K.; Yamamoto, S. Purification and characterization of an acid amidase selective for N-palmitoylethanolamine, a putative endogenous anti-inflammatory substance. J. Biol. Chem. 2001, 276, 35552-35557. 18. Wang, J.; Zhao, L. Y.; Uyama, T.; Tsuboi, K.; Wu, X. X.; Kakehi, Y.; Ueda, N. Expression and secretion of N-acylethanolamine-hydrolysing acid amidase in human prostate cancer cells. J Biochem 2008, 144, 685-90. 19. Ueda, N.; Yamanaka, K.; Yamamoto, S. Purification and Characterization of an Acid Amidase Selective for N-Palmitoylethanolamine, a Putative Endogenous Anti-inflammatory Substance. Journal of Biological Chemistry 2001, 276, 35552-35557. 20. Tai, T.; Tsuboi, K.; Uyama, T.; Masuda, K.; Cravatt, B. F.; Houchi, H.; Ueda, N. Endogenous Molecules Stimulating N-Acylethanolamine-Hydrolyzing Acid Amidase (NAAA). ACS Chem Neurosci 2012, 3, 379-85. 21. Tsuboi, K.; Sun, Y.-X.; Okamoto, Y.; Araki, N.; Tonai, T.; Ueda, N. Molecular Characterization of N-Acylethanolamine-hydrolyzing Acid Amidase, a Novel Member of the Choloylglycine Hydrolase Family with Structural and Functional Similarity to Acid Ceramidase. Journal of Biological Chemistry 2005, 280, 11082-11092. 22. Zhao, L.-Y.; Tsuboi, K.; Okamoto, Y.; Nagahata, S.; Ueda, N. Proteolytic activation and glycosylation of N-acylethanolamine-hydrolyzing acid amidase, a lysosomal enzyme involved in the endocannabinoid metabolism. Biochimica et Biophysica Acta (BBA) - Molecular and Cell Biology of Lipids 2007, 1771, 1397-1405. 23. Ueda, N.; Tsuboi, K.; Uyama, T. N-acylethanolamine metabolism with special reference to N-acylethanolamine-hydrolyzing acid amidase (NAAA). Progress in Lipid Research 2010, 49, 299-315. 24. Wei, B. Q. Q.; Mikkelsen, T. S.; McKinney, M. K.; Lander, E. S.; Cravatt, B. F. A second fatty acid amide hydrolase with variable distribution among placental mammals. Journal of Biological Chemistry 2006, 281, 36569-36578. 25. Vandevoorde, S.; Tsuboi, K.; Ueda, N.; Jonsson, K.-O.; Fowler, C. J.; Lambert, D. M. Esters, Retroesters, and a Retroamide of Palmitic Acid: Pool for the First Selective Inhibitors of N-Palmitoylethanolamine-Selective Acid Amidase. Journal of Medicinal Chemistry 2003, 46, 4373-4376. 26. Saturnino, C.; Petrosino, S.; Ligresti, A.; Palladino, C.; Martino, G. D.; Bisogno, T.; Marzo, V. D. Synthesis and biological evaluation of new potential inhibitors of N-acylethanolamine hydrolyzing acid amidase. Bioorganic & Medicinal Chemistry Letters 2010, 20, 1210-1213. 27. Solorzano, C.; Zhu, C.; Battista, N.; Astarita, G.; Lodola, A.; Rivara, S.; Mor, M.; Russo, R.; Maccarrone, M.; Antonietti, F.; Duranti, A.; Tontini, A.; Cuzzocrea, S.; Tarzia, G.; Piomelli, D. Selective N-acylethanolamine-hydrolyzing acid amidase inhibition reveals a key role for endogenous palmitoylethanolamide in inflammation. Proceedings of the National Academy of Sciences 2009, 106, 20966-20971.

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28. Solorzano, C.; Antonietti, F.; Duranti, A.; Tontini, A.; Rivara, S.; Lodola, A.; Vacondio, F.; Tarzia, G.; Piomelli, D.; Mor, M. Synthesis and Structure−Activity Relationships of N-(2-Oxo-3-oxetanyl)amides as N-Acylethanolamine-hydrolyzing Acid Amidase Inhibitors. Journal of Medicinal Chemistry 2010, 53, 5770-5781. 29. Svíženská, I.; Dubový, P.; Šulcová, A. Cannabinoid receptors 1 and 2 (CB1 and CB2), their distribution, ligands and functional involvement in nervous system structures — A short review. Pharmacology Biochemistry and Behavior 2008, 90, 501-511. 30. Lin, S.; Khanolkar, A. D.; Fen, P.; Goutopoulos, A.; Qin, C.; Papahadjis, D.; Makriyannis, A. Novel Analogues of Arachidonylethanolamide (Anandamide): Affinities for the CB1 and CB2 Cannabinoid Receptors and Metabolic Stability. Journal of Medicinal Chemistry 1998, 41, 5353-5361. 31. Guo, Y.; Abadji, V.; Morse, K. L.; Fournier, D. J.; Li, X.; Makriyannis, A. (-)-1 l-Hydroxy-7’-isothiocyanato-l1’’-,dimethylheptyl-A8-THC:A Novel,High-Affinity Irreversible Probe for the Cannabinoid Receptor in the Brain. Journal of Medicinal Chemistry 1994, 37, 3867-3870. 32. Li, C.; Xu, W.; Vadivel, S. K.; Fan, P.; Makriyannis, A. High Affinity Electrophilic and Photoactivatable Covalent Endocannabinoid Probes for the CB1 receptor. Journal of Medicinal Chemistry 2005, 48, 6423-6429. 33. Picone, R. P.; Khanolkar, A. D.; Xu, W.; Ayotte, L. A.; Thakur, G. A.; Hurst, D. P.; Abood, M. E.; Reggio, P. H.; Fournier, D. J.; Makriyannis, A. (-)-7′-Isothiocyanato-11-hydroxy-1′,1′-dimethylheptylhexahydrocannabinol (AM841), a High-Affinity Electrophilic Ligand, Interacts Covalently with a Cysteine in Helix Six and Activates the CB1 Cannabinoid Receptor. Molecular Pharmacology 2005, 68, 1623-1635. 34. West, J. M.; Zvonok, N.; Whitten, K. M.; Wood, J. T.; Makriyannis, A. Mass Spectrometric Characterization of Human N-Acylethanolamine-hydrolyzing Acid Amidase. Journal of Proteome Research 2011, 11, 972-981. 35. West, J. M.; Zvonok, N.; Whitten, K. M.; Vadivel, S. K.; Bowman, A. L.; Makriyannis, A. Biochemical and Mass Spectrometric Characterization of Human N-Acylethanolamine-Hydrolyzing Acid Amidase Inhibition. PLoS One 2012, 7, e43877. 36. Morse, K. L.; Fournier, D. J.; Li, X.; Grzybowska, J.; Makriyannis, A. A novel electrophilic high affinity irreversible probe for the cannabinoid receptor. Life Sciences 1995, 56, 1957-1962. 37. Long, J. Z.; Nomura, D. K.; Vann, R. E.; Walentiny, D. M.; Booker, L.; Jin, X.; Burston, J. J.; Sim-Selley, L. J.; Lichtman, A. H.; Wiley, J. L.; Cravatt, B. F. Dual blockade of FAAH and MAGL identifies behavioral processes regulated by endocannabinoid crosstalk in vivo. Proceedings of the National Academy of Sciences 2009, 106, 20270–20275. 38. Yang, X.; Birman, V. B. In 1,2,4-Triazolide anion: An active nucleophilic catalyst for ester aminolysis, 2009; American Chemical Society: 2009; pp ORGN-187. 39. West, J. M.; Zvonok, N.; Whitten, K. M.; Wood, J. T.; Makriyannis, A. Mass Spectrometric Characterization of Human N-Acylethanolamine-hydrolyzing Acid Amidase. Journal of Proteome Research 2011, 11, 972-981. 40. Zvonok, N.; Williams, J.; Johnston, M.; Pandarinathan, L.; Janero, D. R.; Li, J.; Krishnan, S. C.; Makriyannis, A. Full Mass Spectrometric Characterization of Human Monoacylglycerol Lipase Generated by Large-Scale Expression and Single-Step Purification. Journal of Proteome Research 2008, 7, 2158-2164.

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CHAPTER 6

FUTURE DIRECTIONS

231

6.1 Novel Endocannabinoid Probes

In Chapter 2, AM9017 was identified as one of the first anandamide based probes with

acceptable hCB2 affinity to conduct LAPS studies. So far, covalent assays of AM9017 have only

been performed on the WT and mutant (C6.47S) on the hCB2. To confirm that C6.47 is the only

amino acid residue of the receptor interacting with AM9017, covalent studies need to be

conducted on mutants of the other cysteine residues. If AM9017 only interacts at TMH6, the

assays should exhibit no loss in covalent binding on all other hCB2 cysteine to serine mutant cell

lines.

While AM9017 was the best covalent probe utilizing the isothiocyanate moiety, we have

also identified AM9069 as a metabolically stable anandamide based probe which also exhibited a

high affinity for the hCB2 receptor. This ligand is a very interesting for covalent studies as it is

one of the best azide based probes and is metabolically resistant to hydrolysis from FAAH. To

expand on these metabolically stable compounds, analogs of AM9069 with an overall length of

19 and 18 carbons could be synthesized (AM9069 has 20 carbons). This may further increase

the covalent binding properties of this type of compound.

Other future research regarding these probes could utilize the terminal alkyne at the head

position for “click” chemistry.1,2 Once the ligand has covalently attached to the receptor an

attempt can be made to fish out the ligand-receptor complex through click chemistry to some

type of linker that is attached to a support. This could aid in the purification and isolation of the

cannabinoid receptors towards an ultimate goal of crystallization.

It would also be interesting to observe the prolonged effect associated with administering

covalent ligands in vivo. Specifically noting the duration of action of a covalent probe compared

to other metabolically stable anandamide analogs.

232

6.2 Chemoenzymatic Methods

While the use of enzymes for chemical transformations is not new, much research can be

applied to chemoenzymatic methods in organic media. Future experiments could focus on

efficient oxidation of the terminal olefin of arachidonic acid or arachidonic acid methyl ester.

This oxidation followed by cleavage could produce a homoallylic triene alcohol that is a

common precursor for tail modified analogs synthesized through the Wittig reaction.3 This

would allow use of arachidonic acid as a starting material, as opposed to synthesizing the

polyolefin from alkyne precursors.

6.3 2-AG Analogs

AM10336 has been identified as a 2-AG analog that is stable in the presence of MGL as

seen in Chapter 4. Future investigation should be focused on finding suitable analogs for the

arachinoate portion of the compound while keeping the dimethylglycerol headgroup. AM10336

so far has been shown to be chemically and metabolically stable. Pharmacological studies on

this ligand should continue, including finding an appropriate model of the effect exhibited by

endogenous 2-AG to compare analogs with.

6.4 NAAA Inhibitors

Investigation of NAAA and identifying classes of inhibitors is still in the early stages.

We have identified AM9053, a stable isothiocyanate inhibitor, as a selective, competitive, and

reversible inhibitor toward NAAA. However, much can be improved on this scaffold, most

notably, reducing the ClogP. As it stands, these compounds are extremely lipophilic with a

ClogP >7. Too improve the drug-ability of these compounds some additional heteroatoms will

need to be included. Possibilities include the introduction of a heterocyclic moiety, such as

imidazole or pyridine, and including an ether linkage in the 10-carbon chain of AM9053. These

233

adjustments can lower the ClogP to the 3-4 range (based on ChemDraw calculations).

Additional scaffolds should be explored; carbamates could be improved to where there is an

acceptable selectivity over FAAH, while increasing the inhibition of NAAA. Concurrently,

AM9053 should be submitted for a pharmacological profile in animal models, and submitted for

studies regarding its influence on inflammation.

6.5 References 1. Kolb, H. C.; Finn, M. G.; Sharpless, K. B. Click Chemistry: Diverse Chemical Function from a Few Good Reactions. Angewandte Chemie. International Ed. In English 2001, 40, 2004-2021. 2. Huisgen, R. Proceedings of the Chemical Society. October 1961. Proceedings of the Chemical Society 1961, 357-396. 3. Yao, F.; Li, C.; Vadivel, S. K.; Bowman, A. L.; Makriyannis, A. Development of novel tail-modified anandamide analogs. Bioorganic & Medicinal Chemistry Letters 2008, 18, 5912-5915.

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APPENDIX I

PUBLICATIONS

Chemoenzymatic synthesis of 2-arachidonoylglycerol, an endogenous ligand forcannabinoid receptors

Subramanian K. Vadivel ⇑, Kyle M. Whitten, A. MakriyannisCenter for Drug Discovery, 116 Mugar Hall, 360 Huntington Avenue, Northeastern University, Boston, MA 02115, USA

a r t i c l e i n f o

Article history:Received 2 December 2010Revised 30 December 2010Accepted 10 January 2011Available online 18 January 2011

Keywords:CannabinoidEndogenous ligand2-ArachidonoylglycerolBiocatalysis

a b s t r a c t

A simple and efficient synthesis of 2-arachidonoyl glycerol, an endogenous agonist for cannabinoid recep-tors was achieved using Novozym 435, immobilized lipase from Candida antarctica.

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2-Arachidonoylglycerol (2-AG) is an endogenous cannabinergicligand that interacts with both CB1 and CB2 receptors. Although 2-AG synthesis involves several candidate enzymes,1 2-AG is inacti-vated principally by monoacylglycerol lipase2(MGL), although fattyacid amide hydrolase (FAAH) may contribute to its degradation.3 2-AG was shown to possess various biological activities, such asbinding to CB1 and CB2 cannabinoid receptors, inhibition of aden-ylyl cyclase in mouse spleen cells, and inducing hypothermia,reducing spontaneous activity, analgesia, and immobility in mice.4

2-AG acts as a full cannabinergic agonist, and the structure of 2-AGis strictly recognized by the cannabinoid receptors (CB1 and CB2).Thus, 2-AG rather than anandamide may represent the true naturalligand for cannabinoid receptors.5–7

The major problem in the synthesis of pure 2-AG is the rapidmigration of the arachidonoyl group from the secondary to the pri-mary hydroxyl group, resulting in the formation of more stable 1-arachidonoyl glycerol. This migration is catalyzed by water, acid,base, or heat.8 Earlier synthetic methods utilized coupling of 1,3-si-lyl9 or benzylidine10 protected glycerol with arachidonic acid andfollowed by deprotection and separation of the isomeric arachido-noyl glycerols. All these methods suffer from extended reactiontime, acidic conditions required for the removal of the protectinggroups as well as extensive work up and purification. Anotherinteresting method also appeared in the literature, which utilizesregioselective transformation of glycidyl arachidonate into 2-

arachidonoyl-1,3-bis(trifluoroacetyl)glycerol followed by thecleavage of trifluroacetyl group with pyridine.11

Searching for an alternative, green, and efficient methodologythat would circumvent these problems during 2-arachidonoylglycerol synthesis, we have developed an enzyme catalyzed effi-cient and highly regioselective synthesis. The advantage of bioca-talysis is that the reactions are carried out at ambienttemperature, nearly neutral pH and the reactions are often highlyregio- and stereoselective.12,13 Enzymes derived from microbialcells can be immobilized and reused for many cycles. It was knownthat enzymes can show selectivity during glycerolysis of fatty acidsas well as transesterification of symmetrical triglycerides.14–18

Herein, we report an improved, practical application of the abovemethodology in the synthesis of 2-arachidonoyl glycerol utilizingunsymmetrical triglyceride.

The synthesis is based on a two-step enzymatic process(Scheme 1). The first involves the synthesis of 1,3-diacyl glycerolusing the method reported by Halldorsson et al.19 Enzymatic acyl-ation of glycerol was carried out in anhydrous dichloromethaneusing vinylbutyrate as an acyl transfer agent. The reaction pro-ceeded smoothly at 0 �C to provide exclusively 1,3-butyroyl glyc-erol (1) with greater than 90% yield.20 The 1,3-diacylglycerol wascoupled with arachidonic acid using 4-dimethylaminopyridine(DMAP) and 1-(3-dimethyl-aminopropyl)-3-ethylcarbodiimidehydrochloride (EDCI) in dichloromethane at room temperaturefor 12 h providing the required triglyceride (2) in 84% yield.21 Tri-glyeride (2) was subjected to immobilized Candida antarctica (Nov-ozym 435) known for its high 1,3-regioselective glycerolysis oftriglycerides. The reaction was facile and afforded the required 2-arachidonoyl glycerol (3) exclusively (67%) and the by-product

0040-4039/$ - see front matter � 2011 Elsevier Ltd. All rights reserved.doi:10.1016/j.tetlet.2011.01.047

⇑ Corresponding author. Address: Center for Drug Discovery, NortheasternUniversity, 360 Huntington Avenue, 116 Mugar LifeSciences Building, Boston, MA02115, USA. Tel.: +1 617 373 7620; fax: +1 617 373 7493.

E-mail address: [email protected] (S.K. Vadivel).

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butylacetate was easily removed under vacuum. We also noted theformation of ethyl arachidonate (27%) as a side product. It is note-worthy that there was no formation of 1-arachidonoylglycerol ob-served in the reaction. The crude product was purified by a smallfilter column of boric acid impregnated silica.22

In summary, the improved and practical synthesis of 2-AG wassuccessfully carried out without any isomerization of the more sta-ble 1-arachidonoyl glycerol. The mild neutral conditions, easy sca-lability, and removal of the volatile byproduct provide addedadvantages to the current method for the synthesis of 2-monoacylglycerides.

Acknowledgment

One of the authors (S.K.V.) acknowledges the financial supportfor this research from NIDA (R03 DA029184-01).

References and notes

1. Di Marzo, V.; Petrosino, S. Curr. Opin. Lipidol. 2007, 18, 129.2. Lambert, D. M.; Fowler, C. J. J. Med. Chem. 2005, 48, 5059.3. Goparaju, S. K.; Ueda, N.; Yamaguchi, H.; Yamamoto, S. FEBS Lett. 1998, 422, 69.4. Mechoulam, R.; Ben-Shabat, S.; Hanus, L.; Ligumsky, M.; Kaminski, N. E.;

Schatz, A. R.; Gopher, A.; Almog, S.; Martin, B. R.; Compton, D. R., et al Biochem.Pharmacol. 1995, 50, 83.

5. Sugiura, T.; Kishimoto, S.; Oka, S.; Gokoh, M. Prog. Lipid. Res. 2006, 45, 405.6. Sugiura, T.; Kodaka, T.; Nakane, S.; Miyashita, T.; Kondo, S.; Suhara, Y.;

Takayama, H.; Waku, K.; Seki, C.; Baba, N.; Ishima, Y. J. Biol. Chem. 1999, 274,2794.

7. Sugiura, T.; Kondo, S.; Kishimoto, S.; Miyashita, T.; Nakane, S.; Kodaka, T.;Suhara, Y.; Takayama, H.; Waku, K. J. Biol. Chem. 2000, 275, 605.

8. Martin, J. B. J. Am. Chem. Soc. 1953, 75, 5483.9. Han, L. N.; Razdan, R. K. Tetrahedron Lett. 1999, 40, 1631.

10. Seltzman, H. H.; Fleming, D. N.; Hawkins, G. D.; Carroll, F. I. Tetrahedron Lett.2000, 41, 3589.

11. Stamatov, S. D.; Stawinski, J. Tetrahedron Lett. 2002, 43, 1759.12. Nakamura, K.; Yamanaka, R.; Matsuda, T.; Harada, T. Tetrahedron: Asymmetry

2003, 14, 2659.13. Matsuda, T.; Yamanaka, R.; Nakamura, K. Tetrahedron: Asymmetry 2009, 20,

513.

14. Monteiro, J. B.; Nascimento, M. G.; Ninow, J. L. Biotechnol. Lett. 2003, 25, 641.15. Pfeffer, J.; Freund, A.; Bel-Rhlid, R.; Hansen, C. E.; Reuss, M.; Schmid, R. D.;

Maurer, S. C. Lipids 2007, 42, 947.16. Damstrup, M. L.; Jensen, T.; Sparso, F. V.; Kiil, S. Z.; Jensen, A. D.; Xu, X. J. Am. Oil

Chem. Soc. 2005, 82, 559.17. Damstrup, M. L.; Jensen, T.; Sparso, F. V.; Kiil, S. Z.; Jensen, A. D.; Xu, X. J. Am. Oil

Chem. Soc. 2006, 83, 27.18. Irimescu, R.; Iwasaki, Y.; Hou, C. T. J. Am. Oil Chem. Soc. 2002, 79, 879.19. Halldorsson, A.; Magnusson, C. D.; Haraldsson, G. G. Tetrahedron 2003, 59,

9101. NMR data for compound 1: 1H NMR (CDCl3, 500 MHz) d 4.12–4.27 (m,4H), 4.03–4.12 (m, 1H), 2.20–2.41 (t, J = 7.4 Hz, 4H), 1.67 (sxt, J = 7.42 Hz, 4H),0.96 (t, J = 7.32 Hz, 6H).

20. Magnusson, C. D.; Haraldsson, C. G. Tetrahedron 2010, 66, 2728.21. Preparation of 2: To a solution of 1,3-dibutanoylglyerol (1) (0.500 g,

2.45 mmol) and arachidonic acid (0.74 g, 2.45 mmol) in anhydrousdichloromethane (10 mL) was added DMAP (2.4 g, 19.6 mmol) and EDCI(1.88 g, 9.8 mmol). The resulting solution was stirred for 24 h. Cold water wasadded to the reaction mixture and washed with 5% HCl. The aqueous layer wasextracted with dichloromethane and the combined organic extracts werewashed with water, brine, and dried over sodium sulfate. The reaction mixturewas subjected to flash chromatography (hexanes/acetone; 7:3) to afford puretriglyceride as colorless oil (1.06 g, 88%). IR (neat) 3012, 2931, 1741, 1167,1094 cm�1; 1H NMR (CDCl3, 500 MHz) d 5.30–5.45 (m, 8H), 5.23–5.30 (m, 1H),4.10–4.37 (m, 4H), 2.76–2.91 (m, 6H), 2.34 (t, J = 7.57 Hz, 2H), 2.30 (t,J = 7.57 Hz, 4H), 2.12 (q, J = 7.16 Hz, 2H), 2.06 (q, J = 7.32 Hz, 2H), 1.68–1.75(m, 2H), 1.60–1.68 (m, 4H), 1.23–1.42 (m, 6H), 0.91–0.99 (m, 6H), 0.84–0.91(m, 3H); 13C NMR (CDCl3, 100 MHz) d 173.3, 172.8 (2C), 130.7, 129.2, 128.9,128.8, 128.5, 128.3, 128.0, 127.7, 69.2, 62.3 (2C), 36.1 (2C), 33.8, 31.7, 29.5,27.4, 26.7, 25.8 (3C), 24.9, 22.8, 18.5 (2C), 14.3, 13.8 (2C).

22. Preparation of 2-arachidonoyl glycerol (3): Triglyceride 2 (0.050 g,0.102 mmol) and anhydrous ethanol (0.7 mL) were stirred at roomtemperature and the reaction was started by the addition of Novozym 435(75 mg) and the reaction mixture was stirred for 1 h and the starting materialwas completely consumed. An additional 50 mg was added and the reactionmixture was stirred for another 1 h. After completion of the reaction theenzyme was filtered off and washed with ether and the solvent was evaporatedand the crude product was purified by plug of boric acid impregnated silica gel(hexanes/acetone; 9:1–3:2) to afford analytically pure 2-AG (0.026 g, 67%) as acolorless oil. IR (neat) 3420, 3012, 2927, 1736, 1456, 1378, 1277, 1152 cm�1;1H NMR (500 MHz, CDCl3) d 5.37–5.49 (m, 8H), 4.93 (p, 5.0 Hz, 1H), 3.84 (t,J = 5.0 Hz, 4H), 2.79–2.88 (m, 6H), 2.39 (t, J = 7.5 Hz, 2H), 2.13 (q, J = 7.0 Hz, 2H),2.06 (q, J = 7.0 Hz, 2H), 2.00 (br t, J = 5.5 Hz, 2H), 1.72 (p, J = 7.2 Hz, 2H), 1.30–1.40 (m, 6H), 0.92 (t, J = 7.2 Hz, 3H); 13C NMR (CDCl3, 100 MHz) d 174.1, 130.7,129.3, 128.9, 128.8, 128.5, 128.3, 128.1, 127.8, 75.2, 62.5 (2C), 33.8, 31.7, 29.6,27.4, 26.7, 25.8, 25.8, 25.8, 24.9, 22.8, 14.3.

HOOH

OHHO

OCO(CH2)2CH3

OCO(CH2)2CH3

ImmobilizedCandida antarctica(Novozym 435)

O

O

EDCI, DMAP

ImmobilizedCandida antarctica (Novozyme 435),Ethanol

O

OOH

OH

OHO

O

OOCO(CH2)2CH3

OCO(CH2)2CH3

2-Arachidonoylglycerol

12

3

90%

84%

67%

Scheme 1.

1150 S. K. Vadivel et al. / Tetrahedron Letters 52 (2011) 1149–1150

Application of chemoenzymatic hydrolysis in the synthesisof 2-monoacylglycerols

Kyle M. Whitten, Alexandros Makriyannis, Subramanian K. Vadivel *

Center for Drug Discovery, 116 Mugar Hall, 360 Huntington Avenue, Northeastern University, Boston, MA 02115, Unites States


Article history:Received 13 March 2012Received in revised form 24 April 2012Accepted 25 April 2012Available online 5 May 2012

Keywords:CannabinoidReceptorEndogenous ligand2-MonoacylglycerolBiocatalysis

a b s t r a c t

The selective biocatalyzed synthesis of 2-monoacylglycerols (2-MAGs) through the use of commerciallyavailable immobilized Candida antarctica (Novozym435) and Rhizomucor miehei is explored. Reactions atroom temperature result in the formation of a 2-MAG and a corresponding ethyl ester of the fatty acidwith immobilized C. antarctica within 2 h with yields ranging from 36% to 83%. Similar reaction condi-tions with immobilized R. miehei yielded exclusively the 2-MAG after 24 h with yields ranging from 37%to 88%. Yields vary on the acyl group at the sn-2 position and choice of enzyme involved.


1. Introduction

2-Monoacylglycerols (2-MAGs) exhibit beneficial emulsifyingproperties that are utilized in the food industry,1,2 and in the ad-ministration of pharmaceuticals.3 The polyunsaturated fatty acid(PUFA) occupying the sn-2 position is important in the influence ofstructured triglycerides absorption and digestion.4 The biologicaleffects of fatty acids released from the metabolism of glycerols andamides, or through ingestion, have also been studied.5e7 One of themore intensively studied 2-MAGs, 2-arachidonoylglycerol (2-AG,10b), is a physiologically important lipid signaling molecule actingas a receptor ligand in the endocannabinoid system. Pharmaco-logical properties of 2-AG include hypotension, neuroprotection,and appetite stimulation.8

2-AG and other 2-MAGs in biological systems are usually inac-tivated/catabolized by the hydrolyzing enzyme monoacylglycerollipase (MAGL) to produce a fatty acid and glycerol.9

The synthesis and study of 2-MAGs is made difficult due to acylmigration from the sn-2 to the sn-1 or -3 position (Scheme 1).10,11

This migration is facile and occurs in the presence of acid, base,heat, and protic solvents.12,13 In the case of 2-AG, acyl migrationrenders 1-AG, which is incapable of binding to the endocannabinoidreceptors.14 Many reported syntheses of 2-MAGs involve multiple

laborious steps with unfavorable reaction conditions and work upsthat may promote the unwanted acyl migration. An earlier 2-MAGsynthesis began with the coupling of the fatty acid to a 1,3-triisopropylsilyl (TIPS) glycerol. The removal of the silyl protectinggroups required 24 h with the addition of acetic acid and tetrabu-tylammonium fluoride.15 Another procedure involved coupling offatty acid with 1,3-benzylideneglycerol and removal of the benzyli-denewith phenylboronic acid. The reaction resulted in the formationof the mixture of the 1,3- and 1,2-phenylboronate esters, which wasseparated and cleaved withmethanol and water.16 A third techniqueutilizes the ring opening of a glycidal ester with trifluoroacetic acidto produce a triacylglycerol. The 2-MAG was then formed aftertreatment of the triacylglycerol methanol and pyridine.17

Application of lipase in the syntheses of 1,3-diacylglycerol and1(3)-rac-monoacyl glycerol has been extensively studied andreviewed.18e26 The selectivity and yield are determined by variousfactors, which include amount of enzyme, solvent, temperature,and the type of lipase used.27,28 Even though, the preparation ofselective 1,3-diacylglycerols has been achieved successfully, it hasbeen a challenging task for the synthesis of 2-acylglycerols mainlydue to over hydrolysis and the acyl migration from sn-2 to sn-1 or

Scheme 1.

* Corresponding author. Tel.: þ1 617 373 7620; fax: þ1 617 373 7493; e-mailaddresses: [email protected], [email protected], [email protected](S.K. Vadivel).

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sn-3 position. Lipase-mediated selective hydrolysis of triglycerideusing 1,3-regiospecific lipases, esterification of fatty acids ortransesterification of fatty esters with glycerol, and the glycerolysisof triglycerides have been documented in the literature.29,30 Iri-mescu et al. reported a successful synthesis of various 2-acylglycerols of fatty acids using regiospecific ethanolysis of sym-metrical triglycerides with immobilized Candida antarctica lipase(Novozym 435).22,31 Even though C. antarctica is not considered asa 1,3-regiospecific enzyme, it has been consistently used for thepreparation of 1,3-acylglycerols and ethanolysis of tri-glycerides.31,32 All existing methods utilize symmetrical (‘AAA’type) triglycerides resulting in the formation of corresponding esteras the byproduct that requires exhaustive purification (Fig. 1).

Encouraged by this literature, we recently reported amethod forthe synthesis 2-AG, which utilizes a structured glyceride, 1,3-dibutyryl-2-arachidonate (‘ABA’ type), as a substrate, for we rea-soned that the anticipated byproduct, ethyl butyrate, can be easilyremoved.33 Benefits of this procedure include reactions at ambienttemperature, neutral pH, and conservative reaction time. Themethod is simple and green, as the lipase can be recycled. Never-theless, a significant amount of ethyl arachidonate formed due toover hydrolysis. Since the reaction is selective and proceededquickly, it has become a valuable tool for the radiolabelled synthesisof 2-AG.34 The following work extends our method to the synthesisof 2-acylglycerols starting from saturated and unsaturated fattyacids, and alkyl and aryl carboxylic acids.

2. Results and discussion

To test the general practicality of our method (Scheme 2), wehave synthesized 2-MAGs from various commercially availablelong-chain carboxylic acids, including those of biological impor-tance. The synthesis began with the enzymatic 1,3-diacyl pro-tection of glycerol by the addition of immobilized C. antarctica(Novozym 435) to glycerol and vinyl butyrate in anhydrous CH2Cl2at 0 �C, resulting in the protected glycerol in quantitative yield.32,35

The 1,3-diacylglycerol was then coupled to various medium andlong-chain acids through 1-(3-dimethyl-aminopropyl)-3-ethylcarbodiimide hydrochloride (EDCI) coupling in a 1:1 mixture

of anhydrous THF/CH2Cl2 along with a catalytic amount of 4-dimethylaminopyridine (DMAP) at 0 �C for 4 h. This generatedthe structured triglyceride (‘ABA’ type) in 67e99% yield.

For the hydrolysis step, Novozym 435 was added to the tri-glyceride in a minimal amount of anhydrous ethanol at roomtemperature. By TLC analysis, it was observed that within 1 h thetriglyceride had been completely consumed, and a mixture of 2-MAG, mono-protected 2-MAG, and ethyl butyrate was generated.There is no formation of ethyl ester of fatty acid observed duringthis period. At this point, additional lipase was added to the mix-ture, which was allowed to stir until all the mono-protected 2-MAGwas consumed (1 h), affording the 2-MAG. Some significantamount of ethyl ester was observed during this period, and theformation of ethyl ester largely depended on the type of carboxylicacid used. Aryl and unsaturated carboxylic acids showed more re-sistant toward over hydrolysis compared to saturated fatty acids.

The separation of 1-MAG and 2-MAG is generally performed onboric acid impregnated TLC plates and silica gel columns.36 Non-impregnated silica TLC plates do not resolve 1- and 2-MAG. Thisseparation is a necessary step for most 2-MAG syntheses due to theunfavorable synthetic conditions used, which result in formation ofconsiderable 1-MAG as well. In contrast, the highly regiospecificand neutral reaction conditions when using the lipase result inminimal or no 1-MAG formation. Although silica gel purificationhas been reported to be an inevitable cause of acyl migration in 2-MAG to 1-MAG,37 we did not observe any migration during columnchromatography with untreated silica gel. The only required stepprior to purification was equilibration of silica gel with hexanes.During chromatography, the highly non-polar ethyl ester byprod-uct eluted with ethyl butyrate, and the 2-MAG was collectedwithout any acyl migration. It was observed that the lipase-catalyzed hydrolysis reactions involving saturated triglycerideshad isolated yields <50%, with the ethyl ester byproduct being themajor product, whereas the unsaturated triglycerides had yields inthe range of 55e75%, and triglycerides containing phenylalkylgroups had yields >80% (Table 1). It should also be noted that therewas no observable difference in rate of reaction or isolated yieldfrom the hydrolysis of a 1,3-diacetylglycerol-protected compoundas compared to the 1,3-dibutrylglycerol-protected compound.

We also screened other 1,3-specific lipases to investigatewhether the transformation can be performed in better yield andselectivity toward the range of substrates and found that lipasefrom Rhizomucor miehei showed excellent selectivity toward hy-drolyzing ‘ABA’ type triglycerides. The reaction proceeded ina similar fashion where the triglyceride was consumed quickly, buthydrolysis of diglycerides took 24e48 h. Even though the reactionproceeded very slowly compared to C. antarctica lipase, the R.miehei lipase offered a remarkable improvement in selectivity,

Fig. 1. Types of triglycerides.

Scheme 2.

K.M. Whitten et al. / Tetrahedron 68 (2012) 5422e5428 5423

providing exclusively 2-acylglycerols in excellent yields withoutformation of ethyl ester byproduct. The saturated and unsaturatedfatty acid triglycerides were hydrolyzed in good yields (75e88%)after 24 h. However, in most of the reactions, some unreacted di-glyceride intermediate remained. Allowing the reaction to proceedfor an additional 24 h or adding more enzyme did not improve theyield. When unreacted diglyceride that was separated from the 2-MAG after 24 h was subjected to an additional treatment of R.miehei lipase the maximal yield once again reached 80% 2-MAGformation. Surprisingly, in contrast to C. antarctica lipase, R. mie-hei lipase showed less reactivity toward aryl esters (13 and 14).

3. Conclusion

Synthesis of 2-MAGs is complicated by the propensity of the acylgroup to shift from the sn-2 to the more stable sn-1 or -3 positions,the acyl migration being promoted by heretofore standard reactionconditions. We demonstrate herein that chemoenzymatic hydro-lysis of structured triglycerides is a mild and efficient means tosynthesize 2-MAGs. The ambient temperature, neutral pH, and lackof caustic work up are conditions, whichmarkedly limit 2-MAG acylmigration. The ability to synthesize 2-MAGs from ‘ABA’ type tri-glycerides is an important aspect of this current methodology in

Table 1Structures and yields of lipase-catalyzed 2-MAGs

Compound no. Triglyceride (a) 2-MAG (b) C. antarcticaa yield (%) R. mieheib yield (%)

2 47 84

3 49 82

4 36 80

5 75 83

6 44 78

7 72 83

8 63 77

9 55 79

10 67 75

11 63 76

12 40 88

13 83 40

14 83 37

a Remaining yield consisted of the ethyl ester of sn-2 acyl group.b Remaining yield consisted of intermediate diglyceride.

K.M. Whitten et al. / Tetrahedron 68 (2012) 5422e54285424

comparison to utilizing ‘AAA’ type triglycerides. An excess of fattyacid is not required for this method, which is important when thepreparation of the modified fatty acid involves laborious multistep-synthesis.38e40 This will further enhance the study of structur-eeactivity relationships of these biologically important lipid sig-naling molecules.

4. Experimental

4.1. General methods

Lipase acrylic resin from C. antarctica and Lipozyme�, immobi-lized from R. miehei were purchased from Sigma Aldrich (USA). Allother reagents were used without prior purification. All reactionswere performed under an atmosphere of argon.

All byproducts, ethyl arachidonate, ethyl butyrate, and all di-glycerides were removed during column chromatography. Allglycerols were purified on a Biotage Isolera One using Luknovaprepackaged 12 g columns equilibrated with hexanes.

Compounds 3ae14a were synthesized following the proceduredescribed for 2a; while compounds 3be14b were synthesized fol-lowing the C. antarctica and R. miehei procedures described for 2b.

4.2. 2-Hydroxypropane-1,3-diyl dibutyrate (1)

Immobilized C. antarctica (750 mg) was added to a solution ofglycerol (2.0 g, 21.6 mmol) and vinyl butyrate (6.2 g, 54.0 mmol) inanhydrous CH2Cl2 (10 mL) at 0 �C. The resulting mixturewas stirredfor 3 h under argon atmosphere. Then, additional lipase (400 mg)was added to the reaction mixture, which was stirred for an addi-tional 2 h at 0 �C. The lipase was filtered off, the solvent wasevaporated off under reduced pressure, and the residue was chro-matographed on silica to yield 1 (5.0 g, 99%) as a colorless oil.Rf¼0.55 (40% ethyl acetate/hexanes). 1H NMR (500 MHz, chloro-form-d) d¼4.20 (dd, J¼11.72, 4.39 Hz, 2H), 4.14 (dd, J¼11.72, 5.86 Hz,2H), 4.04e4.12 (m, 1H), 2.41e2.58 (m, 1H), 2.33 (t, J¼7.32 Hz, 4H),1.67 (sxt, J¼7.32 Hz, 4H), 0.96 (t, J¼7.57 Hz, 4H). The 13C NMRspectral data (100 MHz, CDCl3) are in agreement with literaturevalues.32

4.3. 2-(Dodecanoyloxy)propane-1,3-diyl dibutyrate (2a)

EDCI (383 mg, 2.0 mmol), DMAP (19 mg, 0.16 mmol), and 1(204 mg, 0.88 mmol) were added to a solution of lauric acid(160 mg, 0.80 mmol) in a 1:1 mixture of anhydrous THF/CH2Cl2(10 mL) at 0 �C. The reactionmixturewas allowed to stir for 4 h. Thereaction mixture was then diluted with CH2Cl2 (15 mL) and H2O(15 mL). The organic layer was separated, dried over MgSO4, andthe solvent was removed under reduced pressure. The residue waschromatographed on silica gel (0e15% ethyl acetate/hexanes) toyield 2a (221 mg, 67%) as a colorless oil. Rf¼0.50 (15% ethyl acetate/hexanes). 1H NMR (400 MHz, chloroform-d) d¼5.24e5.31 (m, 1H),4.30 (dd, J¼12.09, 4.03 Hz, 2H), 4.16 (dd, J¼12.46, 5.86 Hz, 2H),2.27e2.35 (m, 6H),1.57e1.70 (m, 6H),1.21e1.36 (m,16H), 0.92e0.98(m, 6H), 0.88 (t, J¼6.60 Hz, 3H). 13C NMR (100 MHz, chloroform-d)d¼173.4 (2C), 173.2, 77.4, 69.1, 62.3 (2C), 36.1 (2C), 34.4, 32.1, 29.8,29.7, 29.6, 29.5, 29.3, 25.1, 22.9, 18.6 (2C), 14.4, 13.9 (2C). IR (neat,cm�1) 2926, 2855, 1742, 1460. HRMS for C23H42O6Na (MNaþ)437.2881. Calcd 437.2879.

4.4. 1,3-Dihydroxypropan-2-yl dodecanoate (2b, utilizingC. antarctica)

Immobilized C. antarctica (Novozym 435, 100 mg) was added toa solution of 2a (100 mg, 0.24 mmol), stirred in anhydrous EtOH(1 mL). After the full consumption of 2a (1 h, TLC monitoring),

additional lipase (100 mg) was added until reaction completionwas observed (1 h). The reaction mixture was diluted with CH2Cl2(3 mL), and the lipase was filtered off. The solvent was removedunder reduced pressure, and the resulting residue was chromato-graphed on silica gel (10e50% acetone/hexanes) to yield 2b (31 mg,47%) as a white solid. Rf¼0.26 (30% acetone/hexanes).Mp¼56e57 �C. 1H NMR (400 MHz, chloroform-d) d¼4.93 (quin,J¼4.76 Hz,1H), 3.84 (br s, 4H), 2.38 (t, J¼7.69 Hz, 2H), 2.08 (br s, 2H),1.58e1.69 (m, 2H), 1.20e1.37 (m, 16H), 0.88 (t, J¼6.60 Hz, 3H). 13CNMR (100 MHz, chloroform-d) d¼174.3, 75.3, 62.8 (2C), 34.6, 32.1,29.8, 29.7, 29.6, 29.5, 29.3, 25.2 (2C), 22.9,14.4. IR (neat, cm�1) 3352,2922, 2856, 1730, 1464. HRMS for C15H30O4Na (MNaþ) 297.2041.Calcd 297.2042.

4.5. 1,3-Dihydroxypropan-2-yl dodecanoate (2b, utilizingR. miehei)

Lipozyme�, immobilized from R. miehei (100 mg) was added toa solution of 2a (100 mg, 0.24 mmol) stirred in anhydrous EtOH(1 mL). The reaction mixture was stirred for 24 h, diluted withCH2Cl2 (3 mL), and the lipase was filtered off. The solvent was re-moved under reduced pressure, and the resulting residue waschromatographed on silica gel (10e50% acetone/hexanes) to yield2b (55 mg, 84%) as an oil. All spectral data was consistent with thatobtained using the procedure with C. antarctica.

4.6. 2-(Tetradecanoyloxy)propane-1,3-diyl dibutyrate (3a)

Yield 347 mg, 99%; colorless oil. Rf¼0.47 (15% ethyl acetate/hexanes). 1H NMR (399 MHz, chloroform-d) d¼5.32e5.22 (m, 1H),4.29 (dd, J¼4.4, 11.7 Hz, 2H), 4.15 (dd, J¼5.9, 11.7 Hz, 2H), 2.35e2.25(m, 6H), 1.70e1.56 (m, 6H), 1.36e1.19 (m, 20H), 0.94 (t, J¼7.3 Hz,6H), 0.87 (t, J¼6.6 Hz, 3H). 13C NMR (100 MHz, chloroform-d)d¼173.4 (2C), 173.2, 69.1, 62.3 (2C), 36.1 (2C), 34.4, 32.2, 29.91, 29.89(2C), 29.86, 29.7, 29.6, 29.5, 29.3, 25.1, 22.9, 18.6 (2C), 14.4, 13.9 (2C).IR (neat, cm�1) 2925, 2854, 1741, 1460. HRMS for C25H46O6Na(MNaþ) 465.3195. Calcd 465.3192.

4.7. 1,3-Dihydroxypropan-2-yl tetradecanoate (3b)

C. antarctica: 34 mg, 49%; R. miehei: 57 mg, 82%; white solid.Rf¼0.22 (30% acetone/hexanes). Mp¼57e58 �C. 1H NMR (399 MHz,chloroform-d) d¼4.93 (quin, J¼4.8 Hz, 1H), 3.89e3.78 (m, 3H), 2.38(t, J¼7.3 Hz, 2H), 2.17e2.10 (m, 2H),1.70e1.58 (m, 2H), 1.38e1.19 (m,20H), 0.88 (t, J¼7.3 Hz, 3H). 13C NMR (100 MHz, chloroform-d)d¼174.3, 75.2, 62.8 (2C), 34.6, 32.2, 29.91, 29.87 (2C), 29.8, 29.7,29.6, 29.5, 29.3, 25.2, 22.9, 14.4. IR (neat, cm�1) 3418, 2926, 2855,1729, 1466. HRMS for C17H34O4Na (MNaþ) 325.2354. Calcd325.2355.

4.8. 2-(Palmitoyloxy)propane-1,3-diyl dibutyrate (4a)

Yield 309 mg, 84%; colorless oil. Rf¼0.39 (15% ethyl acetate/hexanes). 1H NMR (500 MHz, chloroform-d) d¼5.30e5.25 (m, 1H),4.30 (dd, J¼4.2, 12.0 Hz, 2H), 4.16 (dd, J¼5.9, 11.7 Hz, 2H), 2.35e2.27(m, 6H), 1.70e1.58 (m, 6H), 1.34e1.21 (m, 24H), 0.98e0.92 (m, 6H),0.88 (t, J¼6.8 Hz, 3H). 13C NMR (100 MHz, chloroform-d) d¼173.4,173.2 (2C), 69.1, 62.3 (2C), 36.1, 34.4, 32.2, 29.9 (6C), 29.7, 29.6, 29.5,29.3, 25.1 (2C), 18.6 (3C), 14.4, 13.9 (2C). IR (neat, cm�1) 2924, 1742,1460. HRMS for C27H50O6Na (MNaþ) 493.3503. Calcd 493.3505.

4.9. 1,3-Dihydroxypropan-2-yl palmitate (4b)

C. antarctica: 25 mg, 36%; R. miehei: 56 mg, 80%; white solid.Rf¼0.27 (30% acetone/hexanes). Mp¼64e65 �C. 1H NMR (500 MHz,chloroform-d) d¼4.93 (quin, J¼4.8 Hz, 1H), 3.84 (t, J¼4.9 Hz, 4H),


2.38 (t, J¼7.6 Hz, 2H), 2.13e2.05 (m, 2H), 1.69e1.59 (m, 2H),1.38e1.20 (m, 24H), 0.88 (t, J¼7.3 Hz, 3H). 13C NMR (100 MHz,chloroform-d) d¼174.3, 75.2, 62.8 (2C), 34.6, 32.2, 29.93, 29.92,29.89, 29.84, 29.7, 29.6, 29.5, 29.3, 25.2 (2C), 23.3, 22.9, 14.4. IR(neat, cm�1) 3320, 2917, 2850, 1730, 1471. HRMS for C19H38O4Na(MNaþ) 353.2668. Calcd 353.2668.

4.10. (Z)-2-(hexadec-9-enoyloxy)propane-1,3-diyl dibutyrate(5a)

Yield 291 mg, 79%; colorless oil. Rf¼0.50 (15% ethyl acetate/hexanes). 1H NMR (399 MHz, chloroform-d) d¼5.38e5.32 (m, 2H),5.31e5.24 (m, 1H), 4.30 (dd, J¼4.4, 11.7 Hz, 2H), 4.16 (dd, J¼5.9,12.5 Hz, 2H), 2.38e2.26 (m, 6H), 2.01 (q, J¼6.6 Hz, 4H), 1.72e1.56(m, 6H), 1.39e1.20 (m, 16H), 0.95 (t, J¼7.3 Hz, 6H), 0.88 (t, J¼7.0 Hz,3H). 13C NMR (100 MHz, chloroform-d) d¼173.3 (2C), 173.1, 130.2,129.9, 69.1, 62.3 (2C), 36.1 (2C), 34.4, 32.0, 30.0, 29.9, 29.4, 29.3,29.25, 29.21, 27.5, 27.4, 25.1, 22.9, 18.6 (2C), 14.3, 13.8 (2C). IR (neat,cm�1) 3007, 2928, 2856, 1742, 1459. HRMS for C27H48O6Na (MNaþ)491.3347. Calcd 491.3349.

4.11. (Z)-1,3-Dihydroxypropan-2-yl hexadec-9-enoate (5b)

C. antarctica: 46 mg, 66%; R. miehei: 58 mg, 83%; colorless oil.Rf¼0.25 (30% acetone/hexanes). 1H NMR (399 MHz, chloroform-d)d¼5.40e5.31 (m, 2H), 4.92 (quin, J¼4.8 Hz, 1H), 3.86e3.79 (m, 4H),2.37 (t, J¼7.7 Hz, 2H), 2.33 (br s, 2H), 2.05e1.97 (m, 4H), 1.63 (quin,J¼7.3 Hz, 2H), 1.39e1.23 (m, 16H), 0.88 (t, J¼6.6 Hz, 3H). 13C NMR(100 MHz, chloroform-d) d¼174.3, 130.3, 129.9, 75.2, 62.6 (2C), 34.6,32.0, 30.0, 29.9, 29.4, 29.32, 29.30, 29.2, 27.4, 27.4, 25.2, 22.9,14.3. IR(neat, cm�1) 3405, 3008, 2924, 2855, 1736, 1462. HRMS forC19H36O4Na (MNaþ) 351.2512. Calcd 351.2511.

4.12. 2-(Stearoyloxy)propane-1,3-diyl dibutyrate (6a)

Yield 300 mg, 85%; colorless oil. Rf¼0.34 (15% ethyl acetate/hexanes). 1H NMR (500 MHz, chloroform-d) d¼5.30e5.24 (m, 1H),4.30 (dd, J¼4.4, 11.7 Hz, 2H), 4.16 (dd, J¼6.1, 12.0 Hz, 2H), 2.34e2.27(m, 6H), 1.70e1.59 (m, 6H), 1.35e1.20 (m, 28H), 0.98e0.93 (m, 6H),0.88 (t, J¼6.6 Hz, 3H). 13C NMR (100 MHz, chloroform-d) d¼173.4(2C),173.2, 69.1, 62.3 (2C), 36.2 (2C), 34.4, 32.2, 29.9 (4C), 29.89 (3C),29.86, 29.7, 29.6, 29.5, 29.3, 25.1, 22.9, 18.6 (2C), 14.4, 13.9 (2C). IR(neat, cm�1) 2924, 1742, 1460. HRMS for C29H54O6Na (MNaþ)521.3813. Calcd 521.3818.

4.13. 1,3-Dihydroxypropan-2-yl stearate (6b)

C. antarctica: 32 mg, 44%; R. miehei: 56 mg, 78%; white solid.Rf¼0.23 (30% acetone/hexanes). Mp¼68e69 �C. 1H NMR (399 MHz,chloroform-d) d¼4.93 (quin, J¼4.6 Hz, 1H), 3.88e3.82 (m, 4H), 2.38(t, J¼7.7 Hz, 2H), 2.04 (t, J¼5.9 Hz, 2H), 1.65 (quin, J¼7.3 Hz, 2H),1.38e1.19 (m, 28H), 0.88 (t, J¼6.6 Hz, 3H). 13C NMR (100 MHz,chloroform-d) d¼174.3, 75.3, 62.8 (2C), 34.6, 32.2, 29.95 (5C), 29.91(2C), 29.8, 29.7, 29.6, 29.5, 29.3, 25.2, 23.0, 13.8. IR (neat, cm�1)3313, 2916, 2849, 1730, 1472. HRMS for C21H42O4Na (MNaþ)381.2982. Calcd 381.2981.

4.14. (Z)-2-(Oleoyloxy)propane-1,3-diyl dibutyrate (7a)

Yield 290 mg, 82%; colorless oil. Rf¼0.43 (15% ethyl acetate/hexanes). 1H NMR (500 MHz, chloroform-d) d¼5.39e5.30 (m, 2H),5.30e5.24 (m, 1H), 4.30 (dd, J¼4.4, 11.7 Hz, 2H), 4.15 (dd, J¼6.1,12.0 Hz, 2H), 2.36e2.25 (m, 6H), 2.01 (q, J¼6.2 Hz, 4H), 1.71e1.55(m, 6H), 1.38e1.19 (m, 20H), 0.95 (t, J¼7.3 Hz, 3H), 0.88 (t, J¼6.8 Hz,3H). 13C NMR (100 MHz, chloroform-d) d¼173.4 (2C), 173.1, 130.3,129.9, 69.1, 62.3, 36.1 (2C), 34.4, 32.1, 30.0, 29.9, 29.8, 29.6 (2C), 29.4,

29.3, 29.2, 27.5, 27.4, 25.1, 22.9, 18.6 (3C), 14.4, 13.9 (2C). IR (neat,cm�1) 3007, 2925, 1742, 1460. HRMS for C29H52O6Na (MNaþ)519.3658. Calcd 519.3662.

4.15. 1,3-Dihydroxypropan-2-yl oleate (7b)

C. antarctica: 48 mg, 67%; R. miehei: 60 mg, 83%; colorless oil.Rf¼0.30 (30% acetone/hexanes). 1H NMR (399 MHz, chloroform-d)d¼5.41e5.31 (m, 2H), 4.92 (quin, J¼4.8 Hz, 1H), 3.88e3.77 (m, 4H),2.49 (br s, 2H), 2.37 (t, J¼7.7 Hz, 2H), 2.01 (q, J¼6.4 Hz, 4H), 1.63(quin, J¼7.3 Hz, 2H), 1.40e1.19 (m, 20H), 0.88 (t, J¼6.6 Hz, 3H). 13CNMR (100 MHz, chloroform-d) d¼174.4, 130.3, 129.9, 75.1, 62.5 (2C),34.6, 32.1, 30.0, 29.9, 29.8, 29.6, 29.4, 29.33, 29.31, 27.45, 27.38, 25.2(2C), 22.9, 14.4. IR (neat, cm�1) 3415, 3008, 2923, 2854, 1735, 1464.HRMS for C21H40O4Na (MNaþ) 379.2827. Calcd 379.2824.

4.16. 2-((9Z,12Z)-Octadeca-9,12-dienoyloxy)propane-1,3-diyldibutyrate (8a)

Yield 344 mg, 98%; colorless oil. Rf¼0.38 (15% ethyl acetate/hexanes). 1H NMR (399 MHz, chloroform-d) d¼5.43e5.30 (m, 4H),5.29e5.24 (m, 1H), 4.30 (dd, J¼4.4, 11.7 Hz, 2H), 4.15 (dd, J¼5.9,11.7 Hz, 2H), 2.77 (t, J¼6.6 Hz, 2H), 2.36e2.26 (m, 6H), 2.05 (q,J¼6.6 Hz, 4H), 1.72e1.56 (m, 6H), 1.41e1.22 (m, 14H), 0.95 (t,J¼7.7 Hz, 6H), 0.89 (t, J¼7.0 Hz, 3H). 13C NMR (100 MHz, chloro-form-d) d¼173.3 (2C), 173.1, 130.5, 130.2, 128.3, 128.1, 69.1, 62.3 (2C),36.1 (2C), 34.4, 31.8, 29.8, 29.6, 29.4 (2C), 29.3, 29.2, 27.4, 25.8, 25.1,22.8, 18.6 (2C), 14.3, 13.9 (2C). IR (neat, cm�1) 3008, 2929, 2856,1741, 1459. HRMS for C29H50O6Na (MNaþ) 517.3506. Calcd 517.3505.

4.17. (9Z,12Z)-1,3-Dihydroxypropan-2-yl octadeca-9,12-dienoate (8b)

C. antarctica: 45 mg, 63%; R. miehei: 55 mg, 77%; colorless oil.Rf¼0.37 (30% acetone/hexanes). 1H NMR (399 MHz, chloroform-d)d¼5.44e5.30 (m, 4H), 4.93 (quin, J¼4.8 Hz, 1H), 3.89e3.76 (m, 4H),2.77 (t, J¼6.6 Hz, 2H), 2.38 (t, J¼7.3 Hz, 2H), 2.13 (t, J¼6.2 Hz, 2H),2.05 (q, J¼6.8 Hz, 4H), 1.69e1.59 (m, 2H), 1.41e1.23 (m,14H), 0.89 (t,J¼6.6 Hz, 3H). 13C NMR (100 MHz, chloroform-d) d¼174.3, 130.5,130.2, 128.3, 128.1, 75.2, 62.8 (2C), 34.6, 31.8, 29.8, 29.6, 29.4, 29.33,29.30, 27.4, 25.9, 25.2 (2C), 22.8, 14.3. IR (neat, cm�1) 3397, 010,2926, 2855, 1736, 1459. HRMS for C21H38O4Na (MNaþ) 377.2667.Calcd 377.2668.

4.18. (Z)-2-(Icos-11-enoyloxy)propane-1,3-diyl dibutyrate (9a)

Yield 328 mg, 88%; colorless oil. Rf¼0.42 (15% ethyl acetate/hexanes). 1H NMR (399 MHz, chloroform-d) d¼5.38e5.32 (m, 2H),5.30e5.24 (m, 1H), 4.30 (dd, J¼4.4, 11.7 Hz, 2H), 4.16 (dd, J¼5.9,12.5 Hz, 2H), 2.36e2.26 (m, 6H), 2.05e1.97 (m, 4H), 1.71e1.57 (m,6H), 1.27 (br s, 24H), 0.95 (t, J¼7.3 Hz, 6H), 0.88 (t, J¼6.6 Hz, 3H). 13CNMR (100 MHz, chloroform-d) d¼173.3 (2C), 173.1, 130.2, 130.0,69.0, 62.3 (2C), 36.1 (2C), 34.4, 32.1, 30.0 (2C), 29.8, 29.7, 29.55 (2C),29.52, 29.51, 29.3, 27.4 (2C), 25.1 (2C), 22.9, 18.6 (2C), 14.4, 13.9(2C).IR (neat, cm�1) 3008, 2925, 2855, 1742, 1459. HRMS forC31H56O6Na (MNaþ) 547.3978. Calcd 547.3975.

4.19. (Z)-1,3-Dihydroxypropan-2-yl icos-11-enoate (9b)

C. antarctica: 40 mg, 55%; R. miehei, 58 mg, 79%; white solid.Rf¼0.24 (30% acetone/hexanes). Mp¼32e33 �C. 1H NMR (399 MHz,chloroform-d) d¼5.38e5.32 (m, 2H), 4.93 (quin, J¼4.8 Hz, 1H),3.89e3.78 (m, 4H), 2.38 (t, J¼7.3 Hz, 2H), 2.20e2.12 (m, 2H), 2.01 (q,J¼6.6 Hz, 4H), 1.71e1.57 (m, 2H), 1.40e1.18 (m, 24H), 0.88 (t,J¼6.6 Hz, 3H). 13C NMR (100 MHz, chloroform-d) d¼174.3, 130.2,130.0, 75.2, 62.8 (2C), 34.6, 32.1, 30.0 (2C), 29.8, 29.7, 29.7, 29.6 (2C),


29.56, 29.51, 29.3, 27.4, 25.2 (2C), 23.0, 14.4. IR (neat, cm�1) 3405,3008, 2923, 2854, 1737, 1465. HRMS for C23H44O4Na (MNaþ)407.3142. Calcd 407.3137.

4.20. 2-((5Z,8Z,11Z,14Z)-Icosa-5,8,11,14-tetraenoyloxy)-propane-1,3-diyl dibutyrate (10a)

Yield 168 mg, 98%; colorless oil. Rf¼0.36 (30% ethyl acetate/hexanes). The 1H and 13C spectral data (500 and 100 MHz, CDCl3)are in agreement with literature values.33 IR (neat, cm�1) 3012,2931, 1741, 1456. HRMS for C31H50O6Na (MNaþ) 541.3502. Calcd541.3505.

4.21. (5Z,8Z,11Z,14Z)-1,3-Dihydroxypropan-2-yl icosa-5,8,11,14-tetraenoate (10b)

C. antarctica: 48 mg, 67%; R. miehei: 54 mg, 75%; colorless oil.Rf¼0.30 (30% acetone/hexanes). The 1H and 13C spectral data (500and 100 MHz, CDCl3) are in agreement with literature values.33 IR(neat, cm�1) 3420, 2013, 2927, 1736, 1456. HRMS for C23H38O4Na(MNaþ) 401.2677. Calcd 401.2668.

4.22. 2-((4Z,7Z,10Z,13Z,16Z,19Z)-Docosa-4,7,10,13,16,19-hexaenoyloxy)propane-1,3-diyl diacetate (11a)

EDCI (111 mg, 0.58 mmol), DMAP (6 mg, 0.06 mmol), and diac-etin (44 mg, 0.25 mmol) were added to a solution of docosahex-aenoic acid (75 mg, 0.23 mmol) in anhydrous CH2Cl2 (5 mL) at 0 �C.The reaction mixture was allowed to stir for 4 h. Upon completion,the reaction mixture was diluted with CH2Cl2 and H2O. The organiclayer was separated, dried over MgSO4, and removed under re-duced pressure. The resulting residue was chromatographed onsilica gel (0e30% ethyl acetate/hexanes) to yield to 11a (111 mg,99%) as a colorless oil. Rf¼0.55 (30% ethyl acetate/hexanes). 1H NMR(500 MHz, chloroform-d) d¼5.47e5.32 (m,12H), 5.30e5.21 (m,1H),4.29 (dd, J¼4.4, 11.7 Hz, 2H), 4.16 (dd, J¼5.9, 12.2 Hz, 2H), 2.93e2.77(m,10H), 2.40 (d, J¼2.9 Hz, 4H), 2.14e2.02 (m, 6H), 0.98 (t, J¼7.6 Hz,3H). The 13C spectral data (100 MHz, CDCl3) and IR data are inagreement with literature values.32 HRMS for C29H42O6Na (MNaþ)509.2880. Calcd 509.2879.

4.23. (4Z,7Z,10Z,13Z,16Z,19Z)-1,3-Dihydroxypropan-2-yl-docosa-4,7,10,13,16,19-hexaenoate (11b)

C. antarctica: 45 mg, 63%; R. miehei: 63 mg, 76%; colorless oil.Rf¼0.29 (30% acetone/hexanes). 1H NMR (399 MHz, chloroform-d)d¼5.52e5.22 (m, 12H), 4.92 (quin, J¼4.6 Hz, 1H), 3.82 (t, J¼5.1 Hz,4H), 2.91e2.77 (m, 10H), 2.49e2.37 (m, 4H), 2.22 (t, J¼6.2 Hz, 2H),2.07 (quin, J¼7.5 Hz, 2H), 0.97 (t, J¼7.3 Hz, 3H). 13C NMR (100 MHz,chloroform-d) d¼173.5, 132.3 (2C), 129.8 (2C), 128.8, 128.6, 128.5(2C), 128.3 (2C), 128.1, 127.9, 75.4, 62.6 (2C), 34.4, 25.8 (5C), 23.0(2C), 20.8. IR (neat, cm�1) 3401, 3013, 2663, 1736, 1390. HRMS forC25H38O4Na (MNaþ) 425.2666. Calcd 425.2668.

4.24. 2-(Docosanoyloxy)propane-1,3-diyl dibutyrate (12a)

Yield 201 mg, 56%; colorless oil. Rf¼0.48 (15% ethyl acetate/hexanes). Mp¼27e28 �C. 1H NMR (399 MHz, chloroform-d)d¼5.32e5.23 (m, 1H), 4.30 (dd, J¼4.4, 11.7 Hz, 2H), 4.16 (dd, J¼5.9,11.7 Hz, 2H), 2.36e2.26 (m, 6H), 1.71e1.57 (m, 6H), 1.25 (s, 36H),0.95 (t, J¼7.3 Hz, 6H), 0.88 (t, J¼6.6 Hz, 3H). 13C NMR (100 MHz,chloroform-d) d¼173.4 (2C), 173.2, 69.1, 62.3 (2C), 36.0 (2C), 34.4,32.2, 29.95 (9C), 29.91 (2C), 29.88, 29.7, 29.6, 29.5, 29.3, 25.1, 22.9,18.6 (2C), 14.4, 13.9 (2C). IR (neat, cm�1) 2923. 2853, 1742, 1462.HRMS for C33H62O6Na (MNaþ) 577.4446. Calcd 577.4444.

4.25. 1,3-Dihydroxypropan-2-yl docosanoate (12b)

C. antarctica: 32 mg, 40%; R. miehei: 70 mg, 88%; white solid.Rf¼0.27 (30% acetone/hexanes). Mp¼79e80 �C. 1H NMR (399 MHz,chloroform-d) d¼4.93 (quin, J¼4.6 Hz, 1H), 3.88e3.81 (m, 4H), 2.38(t, J¼7.7 Hz, 2H), 2.08 (s, 2H), 1.69e1.59 (m, 2H), 1.38e1.19 (m, 36H),0.88 (t, J¼6.2 Hz, 3H). 13C NMR (100 MHz, chloroform-d) d¼174.3,75.2, 62.8 (2C), 34.6, 32.2, 31.8, 29.94 (7C), 29.90 (2C), 29.8, 29.7,29.6, 29.5, 29.3, 25.2, 22.94, 22.89, 14.4. IR (neat, cm�1) 3313, 297,2850, 1730, 1472. HRMS for C25H50O4Na (MNaþ) 437.3610. Calcd437.3607.

4.26. 2-(3-Phenylpropanoyloxy)propane-1,3-diyl dibutyrate(13a)

Yield 280 mg, 98%; colorless oil. Rf¼0.48 (35% ethyl acetate/hexanes). 1H NMR (399 MHz, chloroform-d) d¼7.33e7.25 (m, 2H),7.24e7.15 (m, 3H), 5.31e5.23 (m, 1H), 4.28 (dd, J¼4.4, 11.7 Hz, 2H),4.13 (dd, J¼5.9, 11.7 Hz, 2H), 2.96 (t, J¼7.7 Hz, 2H), 2.66 (t, J¼8.1 Hz,2H), 2.34e2.24 (m, 6H), 1.64 (sxt, J¼7.3 Hz, 4H), 0.94 (t, J¼7.3 Hz,6H). 13C NMR (100 MHz, chloroform-d) d¼173.3 (2), 172.2, 140.4,128.7 (2), 128.5 (2), 126.6, 69.4, 62.2 (2), 36.1 (2), 35.9, 31.0, 18.6 (2),13.9 (2C). IR (neat, cm�1) 3027, 2966, 2877, 1737, 1455. HRMS forC20H28O6Na (MNaþ) 387.1787. Calcd 387.1784.

4.27. 1,3-Dihydroxypropan-2-yl 3-phenylpropanoate (13b)

C. antarctica: 38 mg, 83%; R. miehei: 8 mg, 40%; white foam.Rf¼0.18 (40% acetone/hexanes). 1H NMR (399 MHz, chloroform-d)d¼7.36e7.28 (m, 2H), 7.26e7.18 (m, 3H), 4.89 (td, J¼4.5, 9.3 Hz, 1H),3.79e3.71 (m, 4H), 3.02e2.96 (m, 2H), 2.77e2.70 (m, 2H), 1.91e1.83(m, 2H). 13C NMR (100 MHz, chloroform-d) d¼175.2, 140.4, 128.8(2), 128.5 (2), 126.7, 75.5, 62.6 (2), 36.1, 31.8. IR (neat, cm�1) 3412,3029, 2935, 2881, 1731, 1454. HRMS for C12H16O4Na (MNaþ)247.0945. Calcd 247.0946.

4.28. 2-(5-Phenylpentanoyloxy)propane-1,3-diyl dibutyrate(14a)

Yield 298 mg, 99%; colorless oil. Rf¼0.63 (35% ethyl acetate/hexanes). 1H NMR (500 MHz, chloroform-d) d¼7.31e7.24 (m, 2H),7.21e7.14 (m, 3H), 5.31e5.23 (m, 1H), 4.30 (dd, J¼4.4, 11.7 Hz, 2H),4.14 (dd, J¼5.9, 11.7 Hz, 2H), 2.63 (t, J¼7.1 Hz, 2H), 2.35 (t, J¼6.8 Hz,2H), 2.29 (t, J¼6.8 Hz, 4H), 1.71e1.58 (m, 8H), 0.94 (t, J¼7.3 Hz, 6H).13C NMR (100 MHz, chloroform-d) d¼173.4 (2C), 172.9, 142.2, 128.6(2C), 128.6 (2C), 126.0, 69.2, 62.3 (2C), 36.1 (2C), 35.8, 34.2, 31.0,24.7, 18.6 (2C), 13.8 (2C). IR (neat, cm�1) 3028, 2965, 2876, 1738,1454. HRMS for C22H32O6Na (MNaþ) 415.2094. Calcd 415.2097.

4.29. 1,3-Dihydroxypropan-2-yl 5-phenylpentanoate (14b)

C. antarctica: 50 mg, 83%; R. miehei: 24 mg, 37%; white foam.Rf¼0.26 (40% acetone/hexanes). 1H NMR (399 MHz, chloroform-d)d¼7.31e7.25 (m, 2H), 7.22e7.13 (m, 3H), 4.92 (td, J¼4.8, 9.5 Hz, 1H),3.86e3.76 (m, 4H), 2.64 (t, J¼7.0 Hz, 2H), 2.41 (t, J¼7.0 Hz, 2H),2.17e2.11 (m, 2H), 1.74e1.60 (m, 4H). 13C NMR (100 MHz, chloro-form-d) d¼174.0, 142.2, 128.6 (4C), 126.1, 75.2, 62.7 (2C), 35.8, 34.4,31.0, 24.7. IR (neat, cm�1) 3414, 3027, 2936, 2882, 1731, 1454. HRMSfor C14H20O4Na (MNaþ) 275.1257. Calcd 275.1259.

Acknowledgements

We thank Dr. David Janero for helpful discussions. We thank Dr.Furong Sun at the School of Chemical Sciences, University of Illinoisat Urbana-Champaign, Urbana, IL for supplying HRMS data. We


would like to acknowledge the financial support for this researchfrom NIDA (R03 DA029184-02).

Supplementary data

1H and 13C NMR spectra for all new compounds reported. Sup-plementary data associated with this article can be found in theonline version, at doi:10.1016/j.tet.2012.04.101.


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http://dx.doi.org/doi:10.1016/j.tet.2012.04.101

Enzymatic synthesis of N-acylethanolamines: direct method for theaminolysis of esters

Kyle M. Whitten, Alexandros Makriyannis, Subramanian K. Vadivel ⇑Center for Drug Discovery, Department of Chemistry and Chemical Biology, 116 Mugar Hall, 360 Huntington Avenue, Northeastern University, Boston, MA 02115, USA


Article history:Received 16 July 2012Revised 7 August 2012Accepted 9 August 2012Available online 19 August 2012

Keywords:N-AcylethanolaminesCandida antarcticaEndogenous ligandAmidationBiocatalysis

a b s t r a c t

Immobilized Candida antarctica (Novozyme 435) catalyzed synthesis of N-acylethanolamines isdescribed. Treatment of methyl esters with lipase and amines yielded the desired amides within 2–24 h with yields ranging from 41% to 98%.


N-Acylethanolamines (NAEs), ethanolamides of various long-chain fatty acids, constitute a class of bioactive lipid moleculesformed from glycerophospholipids through the phosphodiester-ase-transacylation pathway consisting of Ca2+-dependent N-acyltransferase and N-acylphosphatidylethanolamine-hydrolyzingphospholipase D.1,2 Among the NAEs, N-arachidonoylethanol-amine, known as anandamide (Fig. 1), is a physiologically impor-tant lipid signaling molecule acting as a receptor ligand in theendocannabinoid system and has been studied extensively.2

Recently, other NAEs such as palmitoylethanolamine and N-oleoylethanolamine (Fig. 1) also gained much attention due totheir anti-inflammatory and analgesic activities and anorexicactivity, respectively.3

NAEs including anandamide are not stored in the cell but ratherproduced on demand, and their endogenous levels are regulateddirectly by enzymes responsible for their formation and degrada-tion. Anandamide has a relatively rapid onset of action, but is rap-idly hydrolyzed by fatty acid amide hydrolase (FAAH) whichaccounts for its short duration of action. Early studies on struc-ture–activity relationships (SAR) focused on the preparationof various amides of arachidonic acid and established thatamides from chloroethylamine, cyclopropylamine, and R-(2)-aminopropanol showed excellent improvement in their respectiveaffinities to the cannabinoid CB1 receptor while exhibiting en-hanced metabolic stability toward FAAH.4–8 Recently, SAR studieson the modification of the hydrophobic chain have gained more

attention and various analogs with fully saturated fatty acid chainsor alternatively encompassed alkyne moieties were synthesized.Furthermore, our laboratory designed and synthesized high affinitycovalent anandamide probes for the CB1 receptor by introducingeither electrophilic isothiocyanato or a photoactivatable azidogroups at the terminal carbon of the arachidonic acid moiety.9 Wehave also studied the effect of aryl substitutions with variablemethylene linker at the distal end of arachidonic acid.10

All the synthetic schemes use the esters of the substituted fattyacids as a starting point and convert them to the needed amidesusing base mediated conventional hydrolysis of an ester to carbox-ylic acid followed by activation of carboxylic acid with either EDCIor CDI and treatment with various amines to provide the respective

O

NH

OH

O

NH

O

NH

H3C(H2C)14OH

OH

arachidonoylethanolamine(anandamide)

N-palmitoylethanolamine

N-oleoylethanolamine

Figure 1. Structures of N-acylethanolamines.

0040-4039/$ - see front matter � 2012 Elsevier Ltd. All rights reserved.http://dx.doi.org/10.1016/j.tetlet.2012.08.042

⇑ Corresponding author. Tel.: +1 617 373 7620; fax: +1 617 373 7493.E-mail address: [email protected] (S.K. Vadivel).

Tetrahedron Letters 53 (2012) 5753–5755

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amides. In few cases, a protected form of ethanolamine is also usedwhich, however, requires an additional deprotection step.

Several methods have been reported for the direct conversionof esters to amides including Mg(OCH3)2 and CaCl2,11 sodiumcyanide,12metal catalysts,13–15 and Alcalase.16 However, most of

these suffer from incomplete conversion, longer reaction times,and possible functional group instability of the final products un-der the conditions used. Here, we report a highly selective lipasemediated mild conversion of esters to biologically importantamides.

O

O

ImmobilizedCandida antarctica

H2NOH

O

NH

OH

hexanes/diisopropylether

Scheme 1. Amidation of esters with immobilized Candida antarctica.

Table 1Amidation of esters with immobilized Candida antarctica in 1:1 hexanes–diisopropylethera

Entry Ester Amine Amide

Isolated yield (%) Time (h)

1

O

O H2N 85 3

2

O

OH2N

OH 98 24

3

O

O H2NOH

89 2

4

OH

O

O H2NOH 85 24

5

N3

O

O H2N 60 24

6

O

OH2N

41 24

7

O

O H2N 85 3

8O

OH3C(H2C)14

H2N 84 3

9

O

O H2N95 24

10

O

O H2NOH 90 24

11

CO2Me

H2N 91 24

a Candida antarctica (Novozyme 435, 100 mg) and amine (0.24 mmol, 1.2 equiv) were added to a stirred solution of ester (0.20 mmol, 1 equiv) in a 1:1 mixture of hexanesand isopropyl ether (1 mL). The reaction was heated to 45 �C and stirred until completion (TLC monitoring). The reaction was diluted with diethyl ether, filtered, andconcentrated. The resulting residue was chromatographed on silica gel to yield the amide.

5754 K. M. Whitten et al. / Tetrahedron Letters 53 (2012) 5753–5755

Lipases have found wide use as biocatalysts for many chemicaltransformations. Many lipases have been studied for their use inamide formation,17,18 such as, amidation of benzyl esters,19 synthe-sis of acetamides in the presence of ionic liquids,20 and acylation ofamines with acids.21 Most of these methods utilize either carbox-ylic acids or vinyl esters of carboxylic acids as reactants and thereactions require relatively high temperatures. In the kinetic reso-lution of amines, Nechab et al. reported that the reaction condi-tions required 80 �C and 3–10 h to acylate chiral amines withCandida antarctica (CAL) and ethyl acetate.22 The aminolysis oflinoleyl ethyl ester with ethanolamine, catalyzed by CAL, in a sol-vent free system produced the linoleylethanolamide only in 24%yield in 20 h including the presence of the unwanted o-acylationproduct.23 While these examples show the use of lipases for theamidation of esters, there is limited reported work on the use oflipases as a direct method for the synthesis of biologically activeNAEs with regard to functional group sensitivity common in thesynthesis of modified fatty acid moieties. We have thus focusedour efforts on the synthesis of biologically active NAEs with immo-bilized CAL from methyl esters and various amines. Developmentin this area will ameliorate the synthesis of multistep tail-modifiedN-acylethanolamines as well as other biologically important fattyacid amide analogs.

To optimize reaction conditions, we chose methyl arachidonateand cyclopropylamine as reactants and hexane as a solvent. Whencarried out at room temperature in the presence of immobilizedCAL, the reaction proceeded smoothly, but very slow as it required24 h for completion. When heated to 45 �C, reaction completionwas observed in a much improved 3 h. For amines not sufficientlysoluble in n-hexane, the reaction proceeded equally well in a 1:1hexanes–diisopropyl ether mixture (Scheme 1). The results areoutlined in Table 1.

Esters and amines were chosen based on their biologicalimportance. Thus methyl arachidonate was treated withcyclopropyl amine, ethanolamine, and (R)-2-aminopropanol to pro-vide arachidonoylcyclopropylamide (ACPA), anandamide, and R-methanandamide, respectively, in excellent yields. Unprotectedethanolamine was directly used in the preparation of variousethanolamides (2, 3, 4, and 10). When performed with a substitutedfatty acid carrying a terminal hydroxyl group (4) the reactionproceeded smoothly to provide the desired amide. There was noobservable transesterification product in any reactions wherehydroxyl groups were present either in the amine or the fatty acidmoieties. To investigate general applicability of the method, wechose various esters and amines and showed that reactions pro-ceeded within 24 h in good yield. Variation in yield was mainlydependent on the amine used. Primary amines, including benzylicamines, underwent amidation smoothly and in excellent yields after24 h. Conversely, cyclohexylamine exhibited slower reactivity withdecreased yield under the present conditions. Longer reaction timesand increased temperature did not improve the yield significantly.Esters of non-fatty acids (9, 10, and 11) underwent amidation withamines in excellent yields and in all the cases the reaction time ap-peared to be more dependent on the amine used.

In summary, we have demonstrated that CAL can be useful forachieving direct formation of amides from various amines andesters containing skipped polyenes, allyl alcohol, allyl azide,alkyne, and aryl moieties. The method described in this report, issimple, efficient, and environmentally friendly and does not re-quire any protection of other susceptible functional groups. Thistransacylation reaction provides excellent yields and is selective.It may find general utility in the synthesis of amides from the cor-responding esters without requiring prior hydrolysis of the esters,as it can be difficult to synthesize amides directly from esters un-der mild conditions. The method should prove to be useful in thesynthesis of drug intermediates and biologically important naturalproducts.

Acknowledgment

One of the authors (S.K.V.) acknowledges the financial supportfor this research from NIDA (R03 DA029184-02).


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K. M. Whitten et al. / Tetrahedron Letters 53 (2012) 5753–5755 5755

Published: October 31, 2011

r 2011 American Chemical Society 972 dx.doi.org/10.1021/pr200735a | J. Proteome Res. 2012, 11, 972–981

ARTICLE

pubs.acs.org/jpr

Mass Spectrometric Characterization of HumanN-Acylethanolamine-hydrolyzing Acid AmidaseJay M. West, Nikolai Zvonok, Kyle M. Whitten, JodiAnne T. Wood, and Alexandros Makriyannis*

Center for Drug Discovery, Northeastern University, Boston, Massachusetts 02115, United States

’ INTRODUCTION

Interest in the recently cloned lysosomal enzyme NAAA hasincreased of late,1�3 driven in part by studies showing that PEAhas significant anti-inflammatory, analgesic, and neuroprotectiveproperties.4�6 These effects have been traced at least partially toits potent endogenous agonism of the peroxisome proliferator-activated receptor-α (PPAR-α).7,8

The potential of blocking N-acylethanolamines (NAEs) likePEA or N-arachidonlyethanolamine (anandamide) from enzy-matic degradation via inhibition as a strategy for pain treatmenthas received considerable interest of late.3 For example, anotherenzyme known to hydrolyze PEA, fatty acid amide hydrolase(FAAH), primarily hydrolyzes the endocannabinoid ananda-mide, and inhibitors of this enzyme have been the subjects ofclinical trials.9

Although functionally very similar to FAAH in its ability tohydrolyze the biologically significant NAEs anandamide andPEA, NAAA has no homology with it and is most similar inprimary amino acid sequence with acid ceramidase (30% homo-logy and 70% similarity), also a lysosomal enzyme, which hydro-lyzes ceramide to fatty acid and sphingosine.3 Possibly due to thelethal consequences of a rare congenital condition where acidceramidase is nonfunctioning (Farber disease), and the identifi-cation of acid ceramidase several decades ago, acid ceramidase is

better characterized than NAAA at present and appears to bebiochemically very similar.10,11 Both acid ceramidase and NAAAare glycoproteins that undergo removal of an N-terminal signalpeptide after biosynthesis, are believed to be transported by themannose-6-phosphate pathway to the acidic late endosomesand/or lysosomes, and are proteolytically activated by an auto-catalytic step under acidic conditions where the polypeptideis cleaved into two chains;10�12 the shorter chain forms theα-subunit (adopting the nomenclature used for acid ceramidasefor NAAA), and the longer chain forming the β-subunit contain-ing the catalytic nucleophile and N-terminal residue cysteine.The other two residues for both enzymes that make up thecatalytic triad, along with cysteine, are aspartate and arginine.

Like acid ceramidase, NAAA suffers from a complete lack ofthree-dimensional structural information, as no crystal or NMRderived structures with significant homology to either enzymeare available. Although homology models have been published inan attempt to map the active site residues for these enzymes,2,11

they may lack reliability as good structural models because ofsuch low homology (<20% amino acid identity) with the

Received: August 3, 2011

ABSTRACT: N-Acylethanolamine-hydrolyzing acid amidase(NAAA) is a lysosomal enzyme that primarily degrades palmi-toylethanolamine (PEA), a lipid amide that inhibits inflam-matory responses. We developed a HEK293 cell line stablyexpressing the NAAA pro-enzyme (zymogen) and a singlestep chromatographic purification of the protein from the me-dia. Matrix-assisted laser desorption/ionization time-of-flightmass spectrometry MALDI-TOF MS analysis of the zymo-gen (47.7 kDa) treated with peptide-N-glycosidase F (PNGaseF) identified 4 glycosylation sites, and acid cleavage of thezymogen into α- and β-subunits (14.6 and 33.3 kDa) activatedthe enzyme. Size exclusion chromatography estimated the massof the active enzyme as 45( 3 kDa, suggesting formation of anα/β heterodimer. MALDI-TOF MS fingerprinting covered morethan 80% of the amino acid sequence, including the N-terminalpeptides, and evidence for the lack of a disulfide bond between subunits. The significance of the cysteine residues was established by theirselective alkylation resulting in almost complete loss of activity. The purified enzyme was kinetically characterized with PEA and a novelfluorogenic substrate, N-(4-methyl coumarin) palmitamide (PAMCA). The production of sufficient quantities of NAAA and a highthroughput assay could be useful in discovering novel inhibitors and determining the structure and function of this enzyme.

KEYWORDS: endocannabinoid, glycosylation, lysosomal enzyme, mannose-6-phosphate, N-acylethanolamine, N-acylethanolamine-hydrolyzing acid amidase (NAAA), N-terminal nucleophile hydrolase, palmitoylethanolamine (PEA)

973 dx.doi.org/10.1021/pr200735a |J. Proteome Res. 2012, 11, 972–981

Journal of Proteome Research ARTICLE

structures available for other members of the choloylglycinehydrolase family, which is part of the N-terminal nucleophilehydrolase superfamily. One major obstacle in the pursuit of athree-dimensional structure for either of these enzymes is thedifficulty in producing sufficient quantities of highly pure protein.

To produce NAAA in sufficient quantity for an initial proteomic-based study, we used mammalian cells and modified expression andpurification protocols previously developed for acid ceramidase10

and for NAAA transiently expressed in HEK293T cells.12,13 Wehave established a HEK293 mammalian system stably expressingNAAA as a gateway for initial proteomic based enzyme character-ization and for assay development. This robust NAAA expressionsystem can be further used to obtain more detail on the post-translational protein modifications, identification of enzyme inhibi-tors, determination of the mechanisms of their action, and forstructural studies.

’EXPERIMENTAL SECTION

MaterialsStandard laboratory chemicals, buffers, culturemedia andmedia

components were purchased from Sigma-Aldrich (St. Louis, MO)and Fisher Chemical (Pittsburgh, PA). Restriction enzymes, DNAligase, and other molecular biology chemicals were obtained fromNew England Biolabs (Beverly, MA).

Vector ConstructionA full-length cDNA of human NAAA inserted into pcDNA

3.1(+) was kindly provided by Dr. Natsuo Ueda of the Departmentof Biochemistry, Kagawa University School of Medicine, Kagawa,Japan. The forward primer 50-TTAAGCTTGAGCCCGAGCC-30

and the reverse primer 50-TCCTCGAGGATCCTTTCTACTC-GGGTTTCT-30 containing incorporatedHindIII and XhoI restric-tion sites (underlined respectively) necessary for cloning, were usedfor PCRamplification of hNAAAcDNAusingPfuDNApolymerase(Stratagene, La Jolla, CA). The fragment was cleaved with therestriction enzymes HindIII and XhoI and inserted by ligation intothe pcDNA 3.1/myc-His C vector treated with the same enzymes.The ligated construct was transformed into One Shot Top10competent E. coli cells, colonies were screened by PCR for thecorrect insert size and DNA sequence was confirmed.

Stable TransfectionHEK293 wild-type cells (American Type Culture Collec-

tion, Manassas, VA) were cultured at 37 �C in a humidifiedincubator (5% CO2) using Dulbecco’s modified Eagle’s med-ium (DMEM) containing 10% fetal bovine serum (FBS), and1% penicillin-streptomycin (P/S). The day before transfectionapproximately 1� 106 HEK293 cells were split into two 75 cm2

culture plates. The purified plasmid pcDNA 3.1/myc-His Cconstruct containing NAAA (48 μg) was linearized with PvuIand added to 120 μL Lipofectamine 2000, then transferredinto the two culture plates of HEK293 cells according to themanufacturers protocol (Invitrogen). Transfected cells wereselected with 600 μg/mL G418 according to a previouslydetermined sensitivity of non transfected HEK293 cells to thisconcentration of antibiotic,14 and after approximately 14 daysindividual colonies were harvested, passed to new culture flasks,and tested for NAAA activity. These colonies with relativelyhigh enzymatic activity were eventually cryopreserved underliquid nitrogen and used for stable expression of NAAA asdetailed below.

Overexpression and PurificationHEK293 cells stably expressing NAAA (with C-terminal hexa-

histidine tag) were cultured at 37 �C in a humidified incubator(5% CO2) on 500 cm

2 plates in DMEMwith 10% FBS, P/S, and0.6 mg/mL Geneticin to approximately 90% confluency. Thenthe culture medium was exchanged for serum-free DMEM withP/S, 0.6 mg/mL Geneticin, and 10 mM NH4Cl and allowed toincubate for 48 h before harvest of the medium. The harvestedmedium was centrifuged at 1000� g for 10 min to removecontaminating cells and ammonium sulfate was added to 60%saturation in four aliquots at 4 �C over a period of one hour. Theculture medium was then centrifuged at 15000� g for 15 min at4 �C and the pellet resuspended in 2% original volume 40 mMphosphate buffer (pH 6.5), 500 mM NaCl (buffer A), anddialyzed into buffer A with two changes at 4 �C. The dialyzedsolution was centrifuged at 15000� g for 15 min at 4 �C, andincubated with approximately 1mL of Talon affinity resin per mgtotal protein for one hour at 4 �C. Thismixture was centrifuged at300� g for 5 min at 4 �C, and the resin washed twice for 15 minwith ten times the resin volume of buffer A containing 25 mMimidazole. The resin pellet was then transferred to a column andNAAA eluted with buffer A containing 150 mM imidazole. Theeluted fraction was dialyzed using 40 mM phosphate buffer(pH 6.5), 150 mM NaCl and 1 mM EDTA (buffer B) with twochanges at 4 �C. The purified protein concentration was deter-mined by the Bradford dye-binding microassay (Bio-Rad), andwas concentrated with Amicon Ultra-0.5 Centrifugal Filters,10 kDa membrane (Millipore), to approximately 2.5 mg/mL(∼50 μM) and stored at 4 �C.

Buffer Exchange ProcedureThe buffer containingNAAA protein was exchanged to another

buffer by reconcentrating 3 times to original volume after 25 folddilution with exchange buffer using 10 kDa membrane AmiconUltra-0.5 Centrifugal Filters (Millipore).

Converting Zymogen to Active Mature NAAA Enzyme byAcid Treatment

Citrate-phosphate buffer (100 mM), pH 4.5, was added topurifiedNAAA at a 4:1 v/v ratio of buffer to protein solution, andincubated for 2 h at 37 �C. For enzymatic assays the acidifiedNAAA was used directly.

Size Exclusion ChromatographyTo evaluate the molecular weight of the active enzyme, we

performed size exclusion chromatography using a Sephacryl-100column (25 � 1 cm). One-hundred micrograms of purifiedhuman NAAA was acidified, dialyzed into buffer B containing2 mM DTT and concentrated to a volume of 50 μL as describedin the previous sections. The concentrated enzyme was manuallyloaded onto the column, and buffer B with 2 mM DTT was runthrough the column at a flow rate of 0.1 mL/min. The molecularweight was determined under similar conditions according toAndrews,15 using the Bio-Rad Gel Filtration Standards: thyro-globulin (670 kDa), γ-globulin (158 kDa), bovine serum albumin(66 kDa, added to standards), ovalbumin (44 kDa), myoglobin(17 kDa), and Vitamin B12 (1750 Da).

SDS-PAGEProtein samples were denatured at 95 �C for 5 min in

Laemmli buffer, and were resolved in SDS-PAGE using AnykD (Bio-Rad) gels. After staining, a FluorChem ImagingSystem (Alpha Innotech Corp., San Leandro, CA) was usedto photograph the gels.



N-(4-Methyl Coumarin)palmitamide (PAMCA) SynthesisPalmitic acid (50 mg, 0.195 mmol) was dissolved in a 1:1

mixture of anhydrous DMF/THF (5 mL) at 0 �C. This solutionwas treated with 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide(45mg, 0.26mmol), 4-dimethylaminopyridine (48mg, 0.39mmol),and 7-amino-4-methyl-coumarin (45 mg, 0.26 mmol). The solutionwas allowed to stir under argon for 24 h. The reaction mixture wasthen diluted with ether (10 mL), washed with water (5 mL), brine(5mL); and dried overMgSO4. The organic layer was concentratedunder reduced pressure and the resulting residue was chromato-graphed on silica gel (13:7 acetone:hexanes) to yield N-(4-methylcoumarin)palmitamide (49.5 mg, 61%) as a white solid. Mp =184�186 �C. 1H NMR (500 MHz, CHLOROFORM-d) δ ppm7.59 (dd, J = 2.0, 8.8 Hz, 1 H) 7.54 (d, J = 8.79 Hz, 1 H) 7.51(d, J = 1.46 Hz, 1 H) 7.29 (br. s., 1 H) 6.21 (s, 1 H) 2.42 (s, 3 H)2.36 (t, J = 7.57Hz, 1 H) 1.70 - 1.78 (m, 2H) 1.24 - 1.33 (m, 24H)0.88 (t, J = 6.8 Hz, 3 H).

NAAA AssayTo have an assay method more conducive to high-throughput

screening than those published for measuring the NAE hydro-lyzing activity of NAAA,2,16,17 we developed the fluorogenic PEAanalog PAMCA, which is hydrolyzed to fluorescent 7-amino-4-methyl coumarin (AMC) and palmitic acid. The assay procedureis similar to that used in the fluorescence-based assays for FAAHand MGL with each point done in triplicate; to each well of a 96well plate 180 μL of 100 mM citrate-phosphate buffer (pH 4.5)containing 3 mMDTT, 0.1% Triton X-100, 0.05% BSA, 150 mMNaCl (assay buffer used by Tsuboi et al.) with 100 ng of purified,acid treated enzyme was added, followed by 20 μL of a dimethylsulfoxide (DMSO) solution containing PAMCA (final concen-tration ranging from 0.5 to 80 μM for the saturation curveexperiment), and then 10 min incubation on a shaking platform.To analyze the activity of the enzyme fractions after the gelfiltration chromatography 10 μL aliquots were added in the samemanner as above with PAMCA at a final concentration of 5 μM.After shaking, the reaction was allowed to proceed at 37 �C for30 min, with fluorescence readings taken every 10 min at a wave-length of 460 nm (using an excitation wavelength of 360 nm) ona Synergy HT Plate Reader using Gen5 software from Bio-Tek.The enzyme activity was calculated by converting the relativefluorescence units to AMC formed, using a standard curve ofAMC dissolved in 10%DMSO-assay buffer. In order to determineenzymatic activity with the native substrate PEA, the purifiedNAAA was first activated by acid treatment as previously de-scribed. The acid treated NAAA (20 ng) was incubated withvarious concentrations of PEA (ranging from 2.5 to 120 μM) in100 μL 10% DMSO-assay buffer for 30 min at 37 �C; eachconcentration was run in duplicate. Blanks, containing assay bufferwithout enzyme, for each PEA concentration were run simulta-neously in duplicate. The reaction was terminated and extractedwith previously published methods.18�20 In short, a mixture ofacetone and ethanol (2:1, v/v) containing deuterated PEA as theinternal standardwas added to each sample,mixed and centrifugedat 4 �C for 5 min at 16000� g. The supernatants were collectedand evaporated under nitrogen stream. Phosphate buffered saline(pH 7.4), methanol and chloroform (100:100:200 μL) wereadded to each sample followed by vigorous vortexing and cen-trifugation. The bottom chloroform fraction was collected anddried under nitrogen stream. Samples were reconstituted in500 μL ethanol and analyzed using LC�MS/MS according to ourpreviously publishedmethods,18�20 using a TSQQuantumUltra

triple quadrupole mass spectrometer (Thermo Electron, SanJose, CA) with an Agilent 1100 HPLC on the front end (AgilentTechnologies, Wilmington, DE). Separation was achieved usingan Agilent Zorbax SB-CN column (2.1 � 50 mm, 5 μm) withgradient elution using 10 mM ammonium acetate (solvent A,pH 7.3) and 100%methanol (solvent B). Eluted peaks were ionizedvia atmospheric pressure chemical ionization (APCI) and de-tected by their respective selected reaction monitoring (SRM)transitions. Enzyme activity was calculated from the decrease inPEA concentration, comparing the blank samples to the enzymesamples. Michaelis�Menten constants were calculated using proFit software (Quantum Soft, Uetikon am See, Switzerland) and aLevenberg�Marquardt algorithm.

MALDI-TOF MS AnalysisBeforeMS analysis or tryptic digestion of activematureNAAA

the acid buffer was exchanged to 50 mM ammonium bicarbonatebuffer as described earlier. For analysis of the intact proteinmasses, 0.5 μL (1.25 μg) of the protein was mixed with 0.5 μLsinapinic acid matrix solution (5 mg/mL dissolved in 50%acetonitrile, 50% water, and 0.1% trifluoroacetic acid) andspotted onto an Opti-TOF 384-well plate insert. For the trypsindigested protein samples, 0.5 μL (1.25 μg) of the digest wasmixed with 0.5 μL α-cyano-4-hydroxycinnaminic acid matrixsolution (5 mg/mL dissolved in 50% acetonitrile, 50% water, and0.1% trifluoroacetic acid) and spotted onto an Opti-TOF 384-well plate insert. MALDI-TOF MS spectra were acquired on a4800 MALDI TOF/TOF mass spectrometer (Applied Biosys-tems, Foster City, CA) fitted with a 200-Hz solid state UV laser(wavelength 355 nm). Spectra of the intact proteins wereacquired in linear mode, and spectra of the peptides wereacquired in reflectron mode. The conditions used for the MSexperiments and instrument calibration were performed asdescribed by Zvonok et al.21

Deglycosylation of Zymogen and Active Mature NAAAProtein for MALDI TOF MS Analysis

To obtain samples of deglycosylated NAAA compatible withMALDI TOF MS analysis, the purified zymogen (10 μg) wasmixed with 1500 units of PNGase F (New England Biolabs,Ipswich, MA) in 50 mM phosphate buffer, pH 7.5, and incubatedfor 48 h at 37 �C.DeglycosylatedNAAA samples were exchangedto 50mM ammonium bicarbonate buffer as described earlier, andspectra were obtained in linear mode using MALDI TOF MSaverage mass measurement. For trypsin digestion the matureNAAA protein was deglycosylated using PNGase F according tothe manufacturer’s protocol as follows: 10 μg of purified, acidtreated enzyme was exchanged into 50mMphosphate buffer, pH7.5, concentrated to 10 μL, denatured by heating for 10 min at90 �C, followed by addition of nonionic detergent NP-40 (finalconcentration 1%) and 500 units of PNGase F and incubation fortwo hours at 37 �C. After deglycosylation the proteins wereresolved by SDS-PAGE, coomassie stained, and the bands wereexcised for trypsin in-gel digestion as described below.

Tryptic DigestionTen micrograms of purified, acid treated NAAA was ex-

changed to 50 mM ammonium bicarbonate buffer, concentratedto 10 μL, and incubated overnight at 37 �C with MS-gradetrypsin (“Trypsin Gold”, Promega) at a NAAA/trypsin mass tomass ratio of 100:1. For the PNGase treated samples, in-geltrypsin digestion was performed. Protein bands were excisedfrom comassie stained SDS-PAGE gels (10 μg protein loaded



into the lane), washed five times with 1 mL ammonium bicar-bonate buffer with shaking for 10min at 37 �C, washed twice with100% acetonitrile for 5 min at room temperature, followed byremoval of the acetonitrile, and then drying the gel pieces for onehour at room temperature. The gel pieces were rehydrated with20 μL 50 mM ammonium bicarbonate buffer containing anappropriate amount of trypsin and incubated overnight at 37 �C.To test the oxidation state of the cysteines, 20 μg of purified, acidtreated enzyme was exchanged to 50 mM ammonium bicarbo-nate buffer and concentrated to 20 μL. NAAA was split into twosamples and in one an aliquot of N-ethylmaleimide (NEM) wasadded to a final concentration of 10 mM, or nothing, andincubated for one hour at room temperature. To the NEMtreated sample dithiothreitol (DTT) was added to a finalconcentration of 25 mM and incubated at 55 �C for one hour.Then an aliquot of iodoacetamide (IAM) was added to theNEM/DTT treated sample to a final concentration of 50 mMand incubated for one hour at room temperature. The NEM/DTT/IAM treated sample along with the untreated controlsample were digested overnight at 37 �C with trypsin at a 100:1NAAA/trypsin ratio.

’RESULTS AND DISCUSSION

Overexpression, Purification, and Kinetic Characterizationof Recombinant Human NAAA

Our initial attempts to overexpress and purify a sufficient amountof NAAA protein for a mass spectrometric characterization andassay development by transient transfection of HEK293T cells werenot very efficient.However, stable transfection ofHEK293 cellswithNAAA cDNA cloned into the pcDNA 3.1 myc-His vector yieldedsignificantly greater quantities of enzyme. The purification of thezymogen from lysed cells and from the culture media afterammonium chloride treatment stimulated secretion of the lyso-somal proteins from the HEK293 cells, similar to the methodpreviously reported,12,13 were simultaneously pursued. The latter

method13 was optimized and found to provide a greater quantityand quality of the lysosomal NAAA, compared to cell lysate thatcontained along with zymogen all cellular components. Virtuallyall zymogen (95%) was precipitated by addition of ammoniumsulfate to 60% saturation following secretion into media,providing partially enriched samples of NAAA. Because theisoelectric point of the pro-enzyme with a hexa-histidine tagwas estimated to be almost neutral (pI 7.3), mildly acidicconditions (pH 6.5) were chosen to avoid protein precipitationand decrease nonspecific binding of other proteins to the IMACresin. This was deemed important as we found albumin(abundant in the media) nonspecifically binding to the Talon

Figure 2. (A) Linear mode MALDI-TOF mass spectra of purifiedhuman NAAA containing the α-subunit, β-subunit, and pro-enzyme(α+β). (B)Majority of the pro-enzyme was converted into theα- and β-subunits after 2 h acid treatment at 37 �C. (C) Purified enzyme after 48 htreatment with PNGase F at 37 �C under native conditions; intense peakat 34810 m/z is that of PNGase F.

Figure 1. SDS-PAGE analysis of human NAAA purification overex-pressed by HEK293 cells. Ten micrograms total protein was loaded intoeach lane. Lane 1, molecular weight markers; lane 2, proteins from themedia precipitated with 60% ammonium sulfate; lane 3, 25 mMimidazole IMACwash fraction; lane 4, 150mM imidazole IMAC elutionfraction; lane 5, 2 h acid treatment of purified NAAA at 37 �C.



resin (identified by excising the 65 kDa band from SDS-PAGEand performing in-gel tryptic digestion and analysis by MALDI-TOF-MS, data not shown). Extensive washing of the resin withphosphate buffer containing a relatively high concentration ofimidazole (25 mM) removed the rest of albumin, and afterelution with phosphate buffer containing 150 mM imidazole wecalculated an overall 75% yield of the enzyme from the NAAAprotein secreted into the media.

As demonstrated by SDS-PAGE and the MALDI-MS spectrashown in Figure 1 and 2A, the protein obtained was nearly pureandmainly in the proenzyme form, although a significant amountof the α- and β-subunits were present. This is similar to previousresults reported for NAAA and acid ceramidase purified fromthe culture media,10,13 and the ratio of proenzyme to α- andβ-subunits in the purified protein varied somewhat from batch tobatch. Taking into account that conversion of the zymogen intoheterodimer, consisting of α- and β-subunits, was ultimatelynecessary to obtain active enzyme for biochemical studies, we didnot pursue to control or prevent a cleavage of some of thezymogen during purification.

Kinetic characterization of the purified active maturehNAAA enzyme was performed by using an LC-MS methodol-ogy previously reported by our laboratory,18�20 where wemeasured enzymatic activity by determining the decrease inPEA concentration. The purified enzyme displayedMichaelis�Menten kinetics, as shown in Figure 3A, with a Km for PEA of21( 3 μM and a Vmax of 5.4( 0.6 nmol/μg/min, similar to thepreviously reported values of a Km of 35 μM and Vmax of1.8 nmol/μg/min for the NAAA enzyme purified from ratlung.22 The maximal velocity of the enzymatic reaction appears

to be somewhat limited by the relatively low solubility of PEA in10%DMSO aqueous assay buffer, as the specific activity did notincrease above a concentration of approximately 40 μM. Weobserved this sharp leveling off of the activity at higher PEAconcentrations every time we repeated the saturation curveexperiment. To simplify and accelerate NAAA activity de-termination in enzyme assays, particularly the screening ofinhibitors, the novel fluorogenic compound PAMCA wassynthesized and tested as a substrate for enzyme. The enzymedisplayed Michaelis�Menten kinetics for PAMCA hydrolysis,with an observed of Km of 6.2( 0.7 μM, as shown in Figure 3B,indicative that the binding affinity of NAAA for this compound issimilar to or slightly greater than that for PEA. The maximalvelocity was 2 orders of magnitude lower, similar to the slowerrate of hydrolysis observed for FAAH andMGL with fluorogenicsubstrates as compared to the natural substrates.21,23 However,the sensitivity, precision, minimal sample handling, and highthroughput capacity of the fluorescent assay more than make upfor the relatively low rate of catalysis with the fluorogenicsubstrate.

Figure 3. Kinetic studies of purified mature human NAAA enzyme (A)with the native substrate palmitoylethanolamine, performed by usingLC�MS/MS assay and (B) with the fluorogenic substrate N-(4-methylcoumarin)palmitamide in fluorescent assay. Assays were performed at37 �C in pH 4.5 buffer, and data were fit to the Michaelis�Mentenequation using a Levenberg�Marquardt algorithm. Mean values ( SDare shown.

Figure 4. Molecular weight of active mature NAAA estimated by sizeexclusive chromatography using a Sephacryl-100 column. SDS-PAGE ofSephacryl-100 column load and elution fractions; lane 1, molecularweight markers; lane 2, purified NAAA; lane 3, purified NAAA after 2 hacid treatment that was loaded onto column; lanes 4�14, eluted columnfractions 5�15 (A). Relative activity of column fractions measured withfluorogenic assay using 10 μL aliquots of each fraction (B). Peak activityobserved in column fraction 7 (lane 6), corresponding to a calculatedmolecular weight of 45 ( 3 kDa.



Evaluation of the Molecular Weight of the Active MatureEnzyme by Size Exclusion Chromatography

Human acid ceramidase, the closest known homologue toNAAA with 70% similarity and 33% identity in amino acidsequence, is a heterodimer composed of α- and β-subunits;10,11

however, NAAA purified from rat lung was found to have onlythe β-subunit.22 Western blot and immunoprecipitation analysisof FLAG tag purified active and inactive mature hNAAAsuggested that the α- and β-subunits formed a complex.12 Toobtain more evidence confirming the subunit structure of theactive mature hNAAA enzyme we performed size exclusivechromatography studies using a Sephacryl S-100 column cali-brated by running a mixture of protein standards. The purifiedzymogen after cleavage by acid treatment, dialysis, and concen-tration was loaded on calibrated Sephacryl S-100 column andelution fractions were collected. The fractions activities weremeasured with the PAMCA substrate hydrolysis assay, while theprotein composition of the fractions were determined using SDS-PAGE analysis. As shown in lane 3 of Figure 4A, the purifiedzymogen was mostly cleaved, formingα- and β-subunits after theincubation at acidic pH. The highest amount of α- and β-subunitsand peak of hydrolyzing activity was observed in elution fractions

7�8 (lane 6�7 of Figure 4A and B). The molecular weight ofprotein in fractions 7�8 was estimated based on calibratedstandard curve as 45 ( 3 kDa, suggesting that the active matureNAAA enzyme is a heterodimer.

MALDI-TOF-MS Analysis of Zymogen, Its Processing andDeglycosylation

As shown in Figure 2A, the linear mode MALDI-TOF massspectrum of the purified enzyme sample confirmed the previousSDS-PAGE results that NAAA had been purified to near homo-geneity. The majority of purified enzyme was in the zymogenform, with an average mass of 47.7 kDa, however the α- andβ-subunits also were observed (14.5 and 33.2 kDa, respectively).In addition, the relative broadness of the peaks suggests that thepurified enzyme pool is heterogeneous due to the variablemodifications of the oligosaccharide side chains in N-glycosy-lated protein. After incubation of the zymogen under acidicconditions, the linear mode MALDI-TOF mass spectrum shownin Figure 2B confirmed that the enzyme had been nearlycompletely converted into the active form comprising the twomain peaks belonging to the α- and β-subunits with an averagemass of 14.6 and 33.3 kDa, respectively.

Figure 5. MALDI-TOFMS spectra of trypsin digest of (A) mature NAAA and (B) deglycosylated mature NAAA obtained in reflectron mode. (Insets)Zoom scan area 2500�3500 Da obtained in linear mode.



We encountered significant difficulty using both enzymatic andchemical approaches in obtaining a fully deglycosylated sample ofNAAA to determine the molecular weight of the enzyme withoutthe oligosaccharide chains. The glycoamidase PNGase F, thatcleaves between the innermost N-acetylglucosamine (GlcNAc)and asparagine resides in N-glycosylated proteins, is the mosteffective with denatured samples, while heat denaturation of NAAAwithout detergents caused the protein to precipitate. Alternatively,the added SDS kept the protein in solution, however it completelysuppressed MS signal detection. Chemical deglycosylation usingtrifluoromethanesulfonic acid24 treatment of lyophilized NAAAresulted in the loss of the protein. Therefore we used a relativelyhigh concentration of PNGase F and incubated with non denaturedNAAA over extended periods (1�3 days) at 37 �C. After 48 hincubation we observed partial deglycosylation (Figure 2C), whilebeyond this time the NAAA proteins could not be detected byMALDI or by SDS-PAGE. It is likely that NAAA is much morestable in its glycosylated form, similar to previous observations withother N-glycosylated proteins.25�27 The spectrum in Figure 2C,obtained at 48 h incubation with PNGase F, shows peaks represent-ing both the fully de- and glycosylated forms, as well as partiallydeglycosylatedNAAA intermediates. The fully deglycosylated forms

were observed in linear mode MALDI-TOFMS as: α-subunit withan average mass of 10972 Da, which is relatively close to thetheoretical mass of 10966Da, the β-subunit with an averagemass of29659 Da, which is close to the theoretical mass of 29662 Da, andthe pro-enzyme with an average mass of 40593 Da, also close to thetheoreticalmass of 40610Da.Thepartially deglycosylatedβ-subunitand zymogen intermediates, as shown in Figure 2C, are comprisedof groups of two and four individual peaks, respectively. Thedistance between peaks in each group is approximately 1500 Da,corresponding to the incremental removal of the N-linked oligo-saccharide chains from glycosylation sites, which is in agreementwith the typical mass of a high mannose type oligosaccharide that isphosphorylated (∼1500 Da).28 Therefore, there are a total of 4 and2 glycosylation sites in the zymogen and in each of the subunits,respectively. Our result agrees with a previous site-directed muta-genesis study of NAAA that identified 4 actual N-glycosylation sites(Asn37, Asn107, Asn309, and Asn333) out of 6 predicted.12

MALDI-TOF MS Fingerprinting of Tryptic Digest of Glycosy-lated and Deglycosylated Mature NAAA

We performed the trypsin digestion of glycosylated and degly-cosylated mature NAAA and characterized the obtained peptides

Table 1. MALDI-TOF MS Fingerprinting of the Purified Mature Human NAAA Digested with Trypsin

position peptide sequences with mass exceeding 500 Da m/z calculated m/z measured error (ppm)

T1/29�35 SPPAAPR 695.3835 695.3828 �0.99T2/36�47 FNVSLDSVPELR 1376.7056b 1376.698 �5.52T3/48�53 WLPVLR 783.4876 783.4904 3.6

T4/54�61 HYDLDLVR 1030.5316 1030.5302 �1.41T5/62�71 AAMAQVIGDR 1031.5302 1031.5302c 0

T7/75�82 WVHVLIGK 951.5774 951.5658 �12.19T8/83�89 VVLELER 857.5091 857.5087 �0.43T9/90�100 FLPQPFTGEIR 1304.6997 1304.7002 0.38

T10-αa/101�125 GMCDFMNLSLADCLLVNLAYESSVF 2755.2486

T10-βa/126�135 CTSIVAQDSR 1079.515 1079.5109 �3.8T10-β+IAM C(IAM)TSIVAQDSR 1136.5364 1136.533 2.992

T10-β+NEM C(NEM)TSIVAQDSR 1204.5627 1204.5626 0.083

T11/136�142 GHIYHGR 839.4271 839.4293 2.65

T12/143�153 NLDYPFGNVLR 1307.6743 1307.6743 0

T14/155�163 LTVDVQFLK 1062.6194 1062.6024 �15.98T15/164�188 NGQIAFTGTTFIGYVGLWTGQSPHK 2680.3518 2680.3652 5.01

T16/189�196 FTVSGDER 910.4265 910.4551 31.39

T18/199�211 GWWWENAIAALFR 1619.8118 1619.7942 �10.87T20/213�221 HIPVSWLIR 1120.6626 1120.6652 2.29

T21/212�236 ATLSESENFEAAVGK 1552.7489 1552.7358 �8.41T23/240�256 TPLIADVYYIVGGTSPR 1821.9745 1821.9734 �0.61T24/257�263 EGVVITR 773.4516 773.4504 �1.5T26/266�283 DGPADIWPLDPLNGAWFR 2039.9974 2040.0183 10.25

T27/284�296 VETNYDHWKPAPK 1584.7805 1584.7699 �6.69T28/297�300 EDDR 534.2154 534.2266 21.01

T30/302�306 TSAIK 519.3137 519.2904 �44.8T31/307�348 ALNATGQANLSLEALFQILSVVPVYNNLTIYTTVMSAGSPDK 4458.2901

T32/349�352 YMTR 570.2704 570.2725 3.61

T37/363�369 GHPFEQK 842.4155 842.4346 22.67

T38/370�387 LISEEDLNMHTGHHHHHH 2180.9791 2180.9756 �1.6aT10 peptide is split into two peptides belonging to different subunits after acid treatment, which was performed prior to trypsin digest. bCalculatedmolecular weight of the T2 peptide was corrected for conversion of asparagine to aspartic acid after deglycosylation by PNGaseF. cT5 monoizotopicpeak coincides with more abundant the T4 + 1 Da isotope peak.



by MALDI-TOF MS fingerprinting (Figure 5). The majority ofthe tryptic peptides with a mass exceeding 500 Da were identified(Table 1) with good mass accuracy (the error typically less than10 ppm and at most 45 ppm between the calculated andmeasuredmasses). We were unable to directly observe the T5 peptide(1031.5302 m/z), most likely because T4 (1030.5316 m/z) wasthe more prominent peptide and its +1 Da isotope peak coincidedwith monoisotopic mass of T5. Analysis of the expected andobserved isotope distribution curves for the T4 and T5 peptidesrevealed their overlapping and confirmed the presence of bothpeptides in the tryptic digest.

It was previously suggested12 that either the first 27 or 28amino acids represent a signal sequence of nascent NAAAprotein, necessary for the translocation of a pro-enzyme intothe endoplasmic reticulum. We identified in the tryptic digestof mature NAAA an ion at 695.4 m/z that after MS/MSfragmentation was confidently assigned to the sequence SPPAAPR(amino acid 29�35 in prepro-enzyme) corresponding to theT1 peptide (Figure 6A). Therefore, the signal peptidase cleavesprepro-enzyme after the signal peptide between the 28th and29th amino acid residues, resulting in an N-terminal serine inthe α-subunit.

The Cys126 of human NAAA was presumed to be located atthe N-terminus of the β-subunit, similar to Cys131 in purified ratlung NAAA,14and shown to be essential for the proteolyticcleavage of the pro-enzyme.12 The cysteine 126 to alaninemutant was unable to undergo self-proteolysis and convert fromthe pro-enzyme to the active mature form,12 leaving someambiguity of it is importance for activity. Here we showed the

significance of the cysteine residue(s) for enzyme activity by theirselective alkylation in mature NAAA, as alkylation by either IAMor NEM resulted in an almost complete loss of activity. We wereable to detect the 1079.5 m/z ion corresponding to the T10-βpeptide and perform its MS/MS analysis, confirming that theN-terminal residue of the β-subunit of the human enzyme is acysteine (Figure 6B).

In an MS based investigation of acid ceramidase all 6 cysteineswere found to be involved in the formation of 3 disulfide bridges,one linking the two subunits, while the two other are locatedin the β-subunit.10 Unexpectedly, the N-terminal cysteine ofβ-subunit that was supposed to be the catalytic nucleophile isinvolved in formation of an intrasubunit disulfide bridge.10 Thereare a total of three cysteines in the primary amino acid sequenceof humanNAAA (two in the α-subunit, Cys103 and Cys113, andone in the b\β-subunit, Cys126). Thus, disulfide bridge formationbetween two subunits is possible only if Cys126 of β-subunitis involved. To determine the oxidation state of Cys126 weperformed a series of cysteine alkylation and reductionexperiments of mature NAAA followed by trypsin digestionand MALDI TOF MS analysis. The treatment of the enzymewith alkylation reagent (IAM or NEM), followed by reductionwith DTT and exposure to the second alkylation reagent (NEMor IAM) should determine whether the cysteine is in a reduced(labeled with the first reagent) or an oxidized state (labeled afterreduction with the second reagent). We performed the experi-ment by reversing the order in which IAM and NEM were usedto ensure the results were not dependent on the reactivity ofthese reagents. When IAM was the first reagent used in the

Figure 6. MALDI-TOF MS/MS analysis of the NAAA tryptic peptides containing the N-terminal sequences of (A) α- and (B) β-subunits.



treatment of mature NAAA, followed by DTT reduction andNEM alkylation, in the trypsin digest we observed a peptide witha molecular weight of 1136.64 m/z that was not present in thenon-alkylated control. Analysis by MS/MS fragmentation re-vealed this peptide to have the sequence CTSIVAQDSR, withthe carbamidomethylated cysteine (data not shown). We ob-served neither nonalkyated (1079.52 m/z), nor NEM modifiedpeptide (1204.56 m/z) with that sequence. When NEM wasadded first, followed by DTT reduction and IAM treatment, onlythe NEM alkylated peptide (1204.56 m/z) was identified intryptic digested mature NAAA. In addition, as previously notednearly all enzymatic activity was extinguished after alkylation byeither IAM or NEM, strongly suggesting that cysteine 126 at theN-terminus of the β-subunit is the catalytic nucleophile and notinvolved in disulfide bridge formation with α-subunit.

There are four actual N-glycosylation sites in mature NAAAthat should be found after trypsin digest in three predictedN-glycosylated peptides � T2NGlc, T10-αNGlc, and T312NGlc.The group of peaks starting at 2593m/z and ending at 3077m/zwas detected using MALDI TOF analysis (Figure 5A, insert) ofmature NAAA digested with trypsin. In the tryptic digest of thedeglycosylated mature NAAA those peaks are not present, whilewe clearly observe the 1376 m/z T2 peptide (Figure 5B, insert).The difference inmass between peptides in this group and the T2peptide is in the range 1200�1700 Da, close to the average massof the oligosaccharide chains we determined from the linearmode MALDI-TOF MS analysis of the intact mature anddeglycosylated enzyme. The most intense 2593 and 3077 m/zpeaks were subjected MS/MS analysis, however we were unableto obtain good quality fragmentation data to identify thestructure and composition of the oligosaccharide chains attachedto glycosylation site in these peptides. The detailed posttransla-tional mature NAAA characterization is beyond the scope of thisinitial proteomic analysis, and will be undertaken later.

We were unable to observe the two other glycosylated ordeglycosylated T10-α and T31 peptides in the trypsin digest ofmature or PNGase F treated NAAA usingMALDI TOFMS. Thedetection of glycosylated peptides was compromised likely dueto heterogeneous glycosylation producing several glycoforms asis typical inN-glycosylated proteins,29 resulting in signal intensityspreading into a group of peaks and their relatively high masses.In addition, extraction of deglycosylated peptides after in geltrypsin digestion was possibly not efficient enough to observethem in the MALDI TOF MS spectra.

Overall, 26 of the 28 total tryptic peptides with masses higherthan 500 Da, covering more than 80% of the mature NAAAamino acid sequence, were identified using MALDI TOF MSfingerprinting.

’CONCLUSION

We established a HEK293 cell line stably expressing humanNAAA and developed a relatively simple single chromatographicstep purification that together are capable of yielding milligramquantities of pure enzyme. We kinetically characterized thepurified human enzyme, and developed a fluorescent assay basedon a novel fluorogenic substrate PAMCA, that has comparableaffinity to the native substrate PEA and allows for a more efficienthigh throughput screening of enzyme inhibitors. We confirmedthe number of glycosylation sites and determined the molecularweights of the fully glycosylated and deglycosylated forms of theenzyme, along with the approximate molecular weights of the

oligosaccharide chains. Our results suggest that the structure ofthe active mature form of the enzyme is a heterodimer, consistingof a non-covalent complex of the α- and β-subunits. MALDITOF MS fingerprinting covered more than 80% of the matureNAAA amino acid sequence including theN-terminal peptides ofthe α- and β-subunits. The putative catalytic nucleophile, whichis the N-terminal cysteine 126 residue of the β-subunit, wasdemonstrated to be in a reduced state. The loss of virtually allenzymatic activity of mature NAAA treated with IAM or NEMwas correlated with cysteine 126 alkylation, determined byMS/MSanalysis of the tryptic digest. Our work establishes that HEK293NAAA expression and purification can provide adequate enzymequantities for more detailed future proteomic analysis, pharmaco-logical characterization and structural studies.

’AUTHOR INFORMATION

Corresponding Author*Alexandros Makriyannis, Ph.D., Northeastern University Centerfor Drug Discovery, 116 Mugar Life Sciences Building, 360Huntington Avenue, Boston, MA 02115. E-mail: [email protected]. Tel.: 617-373-4200. Fax: 617-373-7493.

’ACKNOWLEDGMENT

We thank Prof. Natsuo Ueda, Prof. Kazuhito Tsuboi, and Dr.Toru Uyama for the kind gift of the full length human NAAAclone, a NAAA antibody, and for the valuable advice provided.We thank Dr. Yazen Jmiean and Mahmoud Nasr for experi-mental assistance. This work was supported by grants DA003801,DA007312, and DA009158 from the National Institutes ofHealth/National Institute on Drug Abuse. The views expressedin this publication do not necessarily reflect the official policies ofthe Department of Health and Human Services; nor doesmention by trade names, commercial practices or organizationsimply endorsement by the U.S. Government.

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Biochemical and Mass Spectrometric Characterization ofHuman N-Acylethanolamine-Hydrolyzing Acid AmidaseInhibitionJay M. West, Nikolai Zvonok, Kyle M. Whitten, Subramanian K. Vadivel, Anna L. Bowman,

Alexandros Makriyannis*

Northeastern University, Center for Drug Discovery, Boston, Massachussetts, United States of America

Abstract

The mechanism of inactivation of human enzyme N-acylethanolamine-hydrolyzing acid amidase (hNAAA), with selectedinhibitors identified in a novel fluorescent based assay developed for characterization of both reversible and irreversibleinhibitors, was investigated kinetically and using matrix-assisted laser desorption/ionization time-of-flight massspectrometry (MALDI-TOF MS). 1-Isothiocyanatopentadecane (AM9023) was found to be a potent, selective and reversiblehNAAA inhibitor, while two others, 5-((biphenyl-4-yl)methyl)-N,N-dimethyl-2H-tetrazole-2-carboxamide (AM6701) and N-Benzyloxycarbonyl-L-serine b-lactone (N-Cbz-serine b-lactone), inhibited hNAAA in a covalent and irreversible manner. MSanalysis of the hNAAA/covalent inhibitor complexes identified modification only of the N-terminal cysteine (Cys126) of theb-subunit, confirming a suggested mechanism of hNAAA inactivation by the b-lactone containing inhibitors. Theseexperiments provide direct evidence of the key role of Cys126 in hNAAA inactivation by different classes of covalentinhibitors, confirming the essential role of cysteine for catalysis and inhibition in this cysteine N-terminal nucleophilehydrolase enzyme. They also provide a methodology for the rapid screening and characterization of large libraries ofcompounds as potential inhibitors of NAAA, and subsequent characterization or their mechanism through MALDI-TOF MSbased bottom up-proteomics.

Citation: West JM, Zvonok N, Whitten KM, Vadivel SK, Bowman AL, et al. (2012) Biochemical and Mass Spectrometric Characterization of Human N-Acylethanolamine-Hydrolyzing Acid Amidase Inhibition. PLoS ONE 7(8): e43877. doi:10.1371/journal.pone.0043877

Editor: Andreas Hofmann, Griffith University, Australia

Received June 25, 2012; Accepted July 30, 2012; Published August 31, 2012

Copyright: � 2012 West et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permitsunrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Funding: This work was supported by grants DA003801, DA007312, and DA009158 from the National Institutes of Health/National Institute on Drug Abuse. Thefunders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Competing Interests: The authors have declared that no competing interests exist.

* E-mail: [email protected]

Introduction

Current pharmacological strategies in drug development

targeting the endocannabinoid system are focused on the discovery

of therapeutic agents that selectively modulate the cannabinergic

signaling for the treatment of human disorders, without being

accompanied by undesirable psychotropic side effects [1,2,3].

Toward this goal, two principal endocannabinoid enzymes, fatty

acid amide hydrolase (FAAH) and monoacylglycerol lipase

(MGL), are promising candidates for drug discovery, by modu-

lating the effects of the two principal endocannabinoids,

anandamide (AEA) and 2-arachidonylglycerol (2-AG), respective-

ly, at the cannabinoid receptors 1 and 2 (CB1 and CB2) [4,5].

Other important bioactive fatty acid amides, such as N-

palmitoylethanolamine (PEA), N-oleoylethanolamine (OEA) and

oleamide (OA), with no or very low affinity for the cannabinoid

receptors, are potential substrates in FAAH and N-acylethanola-

mine-hydrolyzing acid amidase (NAAA) regulated metabolism.

NAAA is a lysosomal enzyme that carries out hydrolysis of fatty

acid amides, with its highest activity against PEA [6]. It has been

suggested that the anti-inflammatory, analgesic, and neuroprotec-

tive properties of PEA are mainly due to the activation of

peroxisome proliferator-activated receptor alpha (PPAR-a), and

also in part by activation of the GPR55 and GPR119 receptors

[7,8,9]. Therefore interest in NAAA as a target for novel

therapeutics has been increasing of late [10,11,12], yet unlike

FAAH and MGL there is a complete lack of direct structural

information on the enzyme. The amino acid sequence of hNAAA

has 34% identity with human acid ceramidase (hAC), another

lysosomal enzyme that is not well characterized, which hydrolyzes

ceramide to sphingosine and free fatty acid [13,14]. A very limited

homology of hNAAA and hAC to conjugated bile acid hydrolase

(CBAH) from Clostridium Perfringens, with an available X-ray

structure [15], has been used to generate homology models for

both proteins [12,16]. All three enzymes belong to the cysteine N-

terminal nucleophile (Ntn) hydrolase superfamily, each with the

highly conserved amino acid catalytic triad of Cys, Arg and Asp

[17,18]. CBAH is a single chain protein whose biological assembly

is a homotetramer [15], whereas hNAAA and hAC after self-

catalyzed cleavage of the zymogen at the catalytic nucleophile

Cys126 and Cys143, respectively, under acidic conditions form a-

and b- subunits [16,19]. The hNAAA mutant with Cys126 to

serine substitution was resistant to self-proteolysis and remained

inactive [19]. The precursor zymogen is most likely inactive but

the active mature form of the enzyme is a heterodimer, consisting

of a non-covalent complex of the a- and b-subunits [19,20]. Site-

directed mutagenesis identified four actual N-glycosylation sites in

hNAAA, which were confirmed by our laboratory in a recent

PLOS ONE | www.plosone.org 1 August 2012 | Volume 7 | Issue 8 | e43877

study using mass spectrometry [19,20]. A putative hNAAA

catalytic triad of amino acid residues Cys126, Arg142 and

Asp145 has been predicted from the highly conserved nature of

these residues in the cysteine Ntn hydrolase superfamily and site-

directed mutagenesis experiments, which also identified Glu195 as

a key determinant of acidic cleavage, and suggested that Asn287

plays an important role in proteolytic zymogen activation [19,21].

Recently a mechanism of catalysis via a zwitterionic N-terminal

cysteine in CBAH was proposed based on computational analyses

of free energy simulations and suggested that NAAA may cleave its

substrates using the same catalytic strategy [22].

Previously we overexpressed hNAAA in stably transfected

HEK293 cells and purified the enzyme in an amount sufficient

for its biochemical and proteomic characterization [20]. We have

further optimized hNAAA expression to obtain a three-fold

increase of hNAAA yield. Additionally we developed novel high

throughput fluorescent inhibitor assays for characterization of both

reversible and irreversible hNAAA inhibitors and identified several

potent inhibitors in our compound libraries. 1-isothiocyanatopen-

tadecane (AM9023) is a potent, selective and reversible hNAAA

inhibitor, while 5-((biphenyl-4-yl)methyl)-N,N-dimethyl-2H-tetra-

zole-2-carboxamide (AM6701) and N-Benzyloxycarbonyl-L-serine

b-lactone (N-Cbz-serine b-lactone) inhibit hNAAA in a covalent

and time-dependent manner. The mechanisms of hNAAA

inactivation by AM9023, AM6701 and N-Cbz-serine b-lactone

were investigated using kinetic and MS experimental approaches.

MALDI-TOF analysis of the tryptic digest of hNAAA treated with

AM6701 or N-Cbz-serine b-lactone inhibitor identified modifica-

tion only for the N-terminal cysteine (Cys126) of the b-subunit.

Materials and Methods

MaterialsStandard laboratory chemicals, buffers, culture media and

media components were purchased from Sigma-Aldrich and

Fisher Chemical. AM6701 was made as previously described [23].

N-Cbz-serine b-lactone was purchased from TCI America.

Synthesis of 1-isothiocyanatopentadecane, AM90231,19-thiocarbonyldipyridin-2(1H)-one (65 mg, 0.28 mmol) was

added to a suspension of 1-aminopentadecane (50 mg, 0.22 mmol)

in 5 mL of anhydrous CH2Cl2. Upon completion (0.5 h) the

reaction was quenched with water and the organic layer was

separated and concentrated. The resulting residue was chromato-

graphed on silica to yield AM9023 (55 mg, 94%) as a colorless oil.1H NMR (500 MHz, CHLOROFORM-d) d 3.51 (t, J = 6.80 Hz,

2H), 1.69 (td, J = 6.84, 15.14 Hz, 2H), 1.36–1.44 (m, 2H), 1.21–

1.34 (m, 22H), 0.88 (t, J = 6.84 Hz, 2H). 13C NMR (126 MHz,

CHLOROFORM-d) d 127.3, 45.3, 32.2, 30.2, 30.0, 29.96, 29.94,

29.92 (2C), 29.8, 29.7, 29.6, 29.1, 26.8, 23.0, 14.4. IR (neat) cm21

2924, 2854, 2185, 2090. HRMS for C16H30NS (M-H+) 268.2111.

Calcd. 268.2099.

hNAAA overexpression and purificationThe hNAAA expression and purification was performed as

previously described, with the exception that ammonium chloride

stimulated secretion of zymogen into media was repeated three

times for the same cells, effectively tripling our hNAAA yield [20].

In brief, HEK293 cells stably expressing human NAAA with a C-

terminal hexa-histidine tag were cultured at 37uC in a humidified

incubator (5% CO2) on 500 cm2 plates in DMEM with 10% FBS,

1% penicillin-streptomycin (P/S), and 0.6 mg/mL Geneticin to

approximately 90% confluency. The FBS containing culture

medium was exchanged for 50 ml (per culture plate) serum-free

DMEM with 1% P/S, 0.6 mg/mL Geneticin, 10 mM NH4Cl,

and allowed to incubate for 48 hours. This step was repeated two

more times at 48 hour intervals, with the medium centrifuged to

remove cells and debris and the proteins were precipitated by

adding ammonium sulfate to 60% saturation. The remainder of

the purification was as previously described [20]. The day that

assays or covalent labeling were performed 100 mM citrate-

phosphate buffer, pH 4.5, was added to purified NAAA at a 4:1 v/

v ratio and incubated for 2 hours at 37uC in order to activate the

enzyme.

Fluorometric assay to determine hNAAA inhibition usingN-(4-methyl coumarin)palmitamide (PAMCA) substrate

We previously described the fluorogenic substrate N-(4-methyl

coumarin)palmitamide (PAMCA), which is hydrolyzed by NAAA

to the fluorescent compound 7-amino-4-methyl coumarin (AMC)

and palmitic acid [20]. For hNAAA inhibition we conducted three

point concentration assays with compounds to determine their

potencies and ranges of enzyme inhibition. Purified activated

NAAA (final concentration of 0.25 mg/mL) was incubated in assay

buffer [20] made up to a total volume of 180 mL, followed by

addition of the compound dissolved in 10 mL DMSO (along with

DMSO neat for the control sample) with the final concentrations

for each compound of 1, 10, and 100 mM, in triplicate on a 96 well

plate. These samples were allowed to incubate for 15 min at room

temperature and then 10 mL of a PAMCA stock solution in

DMSO (final PAMCA concentration 10 mM) was added. After

5 minutes of agitation on a shaking plate, the reaction was allowed

to proceed at 37uC for 30 minutes and enzyme activity was

monitored and calculated as previously described [20].

For compounds that inhibited hNAAA in the range of

IC50,1 mM full inhibition curves using eight different concentra-

tions of inhibitor (8 point assay) were generated. To set up 8 point

fluorescent and radioactive assays for each point, the compound in

45 mL DMSO and purified activated NAAA (final enzyme

concentration of 0.25 mg/mL) in 810 mL of NAAA assay buffer

were incubated for 2 hours in order for the covalent compounds to

reach full inhibition. For the fluorescent assays, 190 mL of each of

the above samples (in triplicate) were transferred to a 96 well plate,

followed by addition of 10 mL of a PAMCA stock solution in

DMSO for a final PAMCA concentration of 10 mM. After

5 minutes of agitation on a shaking plate, the reaction was allowed

to proceed at 37uC for 30 minutes and enzyme activity was

monitored and calculated as previously described [20]. The

complete description of the novel fluorescent 8 point assay to

determine the IC50 value for compounds inhibiting hNAAA

activity will be detailed elsewhere.

Radioactive assay to determine IC50 value for reversibleNAAA inhibition using [1,2 214C]N-palmitoylethanolaminesubstrate

The 8 point radioactive assays, similar to that described by

Saturnino et. al., [10] were performed by taking 95 mL of the

enzyme-compound solution described above and adding a 5 mL

solution of [1,2 214C]N-palmitoylethanolamine (10,000 c.p.m./

sample) in DMSO, with a final PEA concentration of 25 mM. The

reaction was performed for 30 minutes at 37uC and terminated by

addition of 200 mL chloroform/methanol (1:1, v/v). The aqueous

layer containing the [1,2 214C]Ethanolamine was quantified by

reading with a 1450 Microbeta Walluc Trilux Liquid Scintillation

and Luminescence Counter. Inhibition constants were calculated

using pro Fit software (Quantum Soft, Uetikon am See,

Switzerland) and a Levenberg-Marquardt algorithm.

Characterization of Human NAAA Inhibition


Fluorometric assay to determine kinact/Ki value forirreversible hNAAA inhibition

The kinact and KI values for covalent inhibitors were determined

similar to Mileni et. al. [24] The fluorescence-based assays were

performed as above with the following exceptions: fluorescence

readings were initiated immediately (data were collected for

120 minutes at 30 second intervals) after mixing the inhibitor

compound and PAMCA substrate (final concentration of

12.4 mM = 26Km) with NAAA in the assay buffer using a

PerkinElmer Wallac EnVision 2104 Multilabel reader to monitor

the fluorescence. The data for each inhibitor concentration ([I])

were fit to a first order equation (Eq. 1) shown below using

KaleidaGraph (Synergy Software, Reading, PA) in order to

determine kobserved (kobs), where Ft is the fluorescence at time t, F0 is

the fluorescence at t = infinite time, F1 is the total fluorescence

change, and kobs is the first order rate constant for enzyme

inactivation. To determine the inhibitor dissociation constant (KI)

and the first order rate constant for enzyme inactivation at infinite

inhibitor concentration (kinact), the kobs values for each [I] obtained

above were fit to a curve using pro Fit software and a Levenberg-

Marquardt algorithm according to Eq. 2, which simplifies to Eq. 3

at [S] = 26Km as used in this experiment.

Ft~F0{F1e{kobst ð1Þ

kobs~kinact½I�

½I�zKI 1z½S�Km

� � ð2Þ

kobs~kinact½I�½I�z3 KIð Þ

ð3Þ

Trypsin digestion of hNAAA treated with inhibitorsTo 10 mg (0.2 nmol) of purified and activated NAAA in 18 mL

of 100 mM citrate-sodium phosphate buffer (pH 4.5) 2 mL of a

DMSO solution containing 2 nmol of the compound of interest

were added or 2 mL DMSO. The inhibitor and DMSO treated

enzyme solutions were incubated at 37uC for 2 hours and then

desalted prior to digestion. These were desalted by re-concentrat-

ing 3 times to original volume after 25 fold dilution with 50 mM

ammonium bicarbonate buffer, pH 8.0, using 10 kDa membrane

Ultra-0.5 Centrifugal Filters (Millipore). The NAAA samples were

incubated overnight at 37uC with MS-grade trypsin (‘‘Trypsin

Gold’’, Promega) at a NAAA:trypsin mass to mass ratio of 100:1.

The tryptic digested NAAA was analyzed immediately or frozen at

280uC for future analysis.

MALDI-TOF-MS Analysis0.5 mL of the trypsin digested NAAA was mixed with 0.5 mL a-

cyano-4-hydroxycinnaminic acid matrix solution (5 mg/mL dis-

solved in 50% acetonitrile, 50% water, and 0.1% trifluoroacetic

acid) and spotted onto an Opti-TOF 384-well plate insert.

MALDI-TOF MS spectra were acquired on a 4800 MALDI

TOF/TOF mass spectrometer (Applied Biosystems, Foster City,

CA) fitted with a 200-Hz solid state UV laser (wavelength

355 nm). The spectra of the peptides were acquired in reflectron

mode. The conditions used for the MS experiments and

instrument calibration were performed as described by Zvonok

et. al. [25]

Molecular ModelingThe sequence for human NAAA was taken from the SWISS-

PROT protein sequence database (amino acids 126–359 primary

accession number Q02083). The homology model of hNAAA was

constructed using the crystal structure of conjugated bile acid

hydrolase (CBAH) from Clostridium Perfringens (PDB ID: 2BJF) [15]

as a template in Prime (1.6 ed., Schrodinger, LLC, New York,

NY). An initial BLAST alignment between the two sequences was

adjusted by taking secondary structure into account using SSpro

and PSIPRED [26]. This alignment was further refined manually

to superimpose Asn204 and Asn287 of hNAAA with Asn82 and

Asn175 of CBAH respectively as previously suggested [12]. The

resultant alignment (13% identity, 21% homology, 34% gaps) was

used for construction of the initial hNAAA model. Loops 2–6 and

8–14 were refined using an ab initio loop prediction algorithm. The

loop refinement step deletes the loop and reconstructs it from a

backbone dihedral library; the loop is then exhaustively sampled to

identify the lowest energy conformation. All other loops featured

mainly homologous residues and contained no gaps or insertions.

The protein underwent a truncated-Newton energy minimization,

using the OPLS_2005 all-atom force field and a Generalized Born

continuum solvation model.

AM6701 and N-Cbz-serine b-lactone were prepared for docking

using the LipPrep (2.2 ed., Schrodinger, LLC, New York, NY)

protocol and the OPLS_2005 force field. The ligands were docked

to hNAAA using the extra precision (XP) procedure in Glide (5.6

ed., Schrodinger, LLC, New York, NY). The top pose for each

ligand was then used to create the product for reaction. A covalent

bond was imposed between the carbonyl carbon of the ligand and

the sulfur atom of Cys126, for AM6701 the leaving group was

removed and for N-Cbz-serine b-lactone the ring was opened.

Atom types were reassigned and the entire system underwent

minimization.

Results and Discussion

hNAAA overexpression and purificationThe multiple harvesting of media containing the secreted

enzyme, after the stimulation of lysosomal protein secretion via

ammonium chloride treatment of the HEK293 cells, was used to

increase the yield of overexpressed hNAAA. With this modifica-

tion the stably transfected HEK293 cells with hNAAA construct

produces ,1 mg of IMAC purified enzyme per 56103 cm2

culture plate area (106500 cm2 culture plates). This amount of

enzyme is sufficient for 26104 data points in fluorescence-based

inhibition assays (,200696 well plates) with 50 ng enzyme per

well, enough to generate full inhibition curves for approximately

800 compounds. All other steps of hNAAA purification were

similar to previously described.16

Kinetic analysis of hNAAA inhibition by AM9023,AM6701, and N-Cbz-serine b-lactone

We previously introduced the novel fluorogenic compound N-

(4-methyl coumarin) palmitamide (PAMCA), which has an affinity

for hNAAA comparable to the native substrate PEA (Km 6.2 mM

and 21 mM for PAMCA and PEA, respectively), and which is

enzymatically hydrolyzed to the fluorescent 7-amino-4-methyl

coumarin (AMC) and palmitic acid [20]. Although the rate of

PAMCA versus PEA hydrolysis is two orders of magnitude slower

the sensitivity, set up time, safety, and rapid readout of the

fluorescence assay makes it superior to the radioactivity based



assay methods. Therefore, PAMCA was selected as a substrate to

develop a high throughput fluorescent inhibition assay to discover

novel hNAAA inhibitors, similar to assays with FAAH and MGL

enzymes [25,27]. We first performed 3 point assay screens of our

compound library to identify potential inhibitors of PAMCA

hydrolysis by hNAAA. The enzyme and compounds at concen-

trations of 1, 10 and 100 mM (3 point assays) were pre-incubated

for 15 min followed by addition of the substrate PAMCA and then

monitoring the increase in fluorescence. For selected compounds

we performed 8 point assays, shown in Figure 1, to obtain full

inhibition curves and IC50 values. AM9023, AM6701 and N-Cbz-

serine b-lactone, identified in the 3 point assay, were characterized

both in radioactive (14C labeled PEA) and fluorescent 8 point

assays to validate the observed IC50 values. The IC50 values for

each selected inhibitor are very similar between those two assays

and are presented in Table 1.

hNAAA inhibition by AM9023AM9023 contains an isothiocyanate group (Figure 2a), and it

was expected that it would react irreversibly with the cysteine

nucleophile in the active site. We have used isothiocyanate based

probes extensively to characterize the cannabinoid receptors,

which covalently react with the cysteines in the receptor

[28,29,30]. However, the following evidence suggested that

AM9023 is a competitive reversible inhibitor: a) the IC50 was

unaffected by longer pre-incubation of hNAAA with AM9023; b)

enzyme activity was fully recovered in a rapid dilution experiment;

c) the profile of a Lineweaver-Burk (double reciprocal) plot

suggested it was a competitive type inhibitor. Yet AM9023 was a

relatively potent inhibitor, with an IC50 = 350670 nM as mea-

sured with the fluorescent assay and an IC50 = 6006130 nM

obtained with the radioactivity based assay (Figures 1a and 1b and

Table 1). AM9023 was selective for hNAAA as compared to

hMGL or rFAAH, which each had an IC50.10 mM. These results

suggested that AM9023 is a reversible and non-covalent inhibitor

of NAAA, and mass spectrometric analysis was used to test this

hypothesis in the mass spectrometric experimental section.

hNAAA inhibition by AM6701AM6701 was recently found to be a very potent inhibitor of

both human MGL and FAAH, with both enzymes having

equivalent IC50 values of 1.2 nM [31]. We identified AM6701

as also a very potent inhibitor of hNAAA with the IC50 values

determined to be 7.262.5 nM (radioactive assay) and

7.760.2 nM (fluorescent assay) as shown in Figures 1a and 1b

and Table 1. The AM6701 potency to inhibit hNAAA was time

dependent; the longer enzyme was preincubated with inhibitor the

lower IC50 was observed (extension of preincubation time to

120 min decreased the IC50 more than 50 fold; data not

presented). A rapid dilution assay experiment with hNAAA and

AM6701 was performed to determine if the inhibition was

reversible. AM6701 was incubated with hNAAA at a concentra-

tion of 100 nM for 120 min, and after twenty-fold dilution to a

final inhibitor concentration of 5 nM there was no recovery of

enzyme activity, even after 24 h incubation, indicating that

inhibition was irreversible, unlike the reversible inactivation of

hMGL by AM6701 [23,32].

hNAAA inhibition by N-Cbz-serine b-lactoneAnother compound, N-Cbz-serine b-lactone, initially discovered

as a hepatitis A virus 3C proteinase inhibitor [33], and more

recently identified as an inhibitor of rat NAAA with an IC50 in the

low micro molar range, was characterized both in radioactive and

fluorescent assays. Structure-activity relationship experiments

suggested that the b-lactone portion of the compound was

essential for inhibition [11,12]. N-Cbz-serine b-lactone inhibited

hydrolysis of the substrate heptadecenoylethanolamide by

HEK293 cell lysate, containing overexpressed recombinant rat

NAAA, with an IC50 = 2.9660.3 mM as determined in LC/MS

based assay [11,12]. The IC50 inhibition values we obtained for

purified hNAAA with the same inhibitor using the radioactive

(IC50 = 1.960.5 mM) and fluorescent (IC50 = 1.760.2 mM) assays,

as shown in Table 1, were similar. The potency of N-Cbz-serine b-

lactone hNAAA inhibition was time dependent; however we

observed a slow regeneration of enzyme catalytic activity in a rapid

dilution experiment of hNAAA inhibited with this compound,

similar to the previously reported partial recovery of initial enzyme

activity (4462%) following 12 h dialysis of rNAAA inhibited with

N-Cbz-serine b-lactone analog.12 The recovery of enzyme activity

after rapid dilution suggested that N-Cbz-serine b-lactone either

introduces enzyme modification(s) that are reversible under assay

conditions or it is a tight-binding reversible inhibitor. It was

proposed that the b-lactone class of inhibitors inhibit NAAA via a

covalent mechanism, that either alkylates or acylates the catalytic

nucleophile cysteine via b-lactone ring opening (Figure 2c).

Figure 1. Concentration dependent inhibition of purifiedhNAAA by three compounds. hNAAA was incubated with thecompounds AM6701 (squares), N-Cbz-serine b-lactone (circles), andAM9023 (diamonds) for two hours in order to reach full inhibitionbefore measuring activity. Panel (A). A radioactivity-based assay with[14C] PEA as substrate. Panel (B). A fluorescence-based assay withPAMCA as substrate. Representative curves are displayed.doi:10.1371/journal.pone.0043877.g001



Characterization of irreversible AM6701 and N-Cbz-serineb-lactone inhibitors of hNAAA

Variable pre-incubation time of the enzyme with the irreversible

inhibitors AM6701 or N-Cbz-serine b-lactone resulted in a

significant variation of IC50. To characterize this class of inhibitors

the use of a second order rate constant, derived from the ratio of

kinact/Ki, has been suggested as the appropriate way to describe

inhibitor potency [24,34]. Unlike IC50 values, kinact/Ki are

independent of pre-incubation times and therefore are a better

measure of potency for irreversible inhibitors [24,34]. To

accurately determine kinact/Ki values it is necessary to follow the

enzymatic reaction continuously, determining the concentration of

either the substrate or product in the course of inhibition, which

we can do with our fluorescence-based assay by monitoring the

fluorescence at 460 nm, the emission peak of the product AMC.

AM6701 was found to have a relatively high kinact/KI of

55000 M21 s21 (Table 1), indicating that AM6701 is a very

potent inhibitor. N-Cbz-serine b-lactone was much less potent

with a kinact/KI of 290 M21 s21. In summary, this novel

fluorescent inhibition assay may be used for characterization of

both reversible (based on IC50 values) and irreversible (based on

kinact/KI values) hNAAA inhibitors.

MALDI-TOF MS analysis of hNAAA inhibition by AM9023,AM6701, and N-Cbz-serine b-lactone

To determine if the selected compounds covalently modified the

enzyme, we employed LAPS methodology similar to that

previously used with hMGL [23]. This approach consists of

incubating the purified enzyme with and without a putative

covalent inhibitor, evaluating extent of inactivation, performing a

tryptic digest, comparing the peptide profile fingerprints using

MALDI-TOF MS, and then assigning the site and nature of any

covalent modification by MS/MS analysis.

MALDI-TOF MS analysis of hNAAA inhibition by AM9023The MALDI-TOF MS spectra of the tryptic digest of untreated

and AM9023 treated hNAAA were identical (data not presented).

This evidence along with the kinetic experiments strongly suggests

that this isothiocyanate based compound is a reversible and non-

covalent inhibitor of hNAAA.

MALDI-TOF MS analysis of hNAAA inhibition by AM6701MALDI-TOF MS analysis of the tryptic digests of untreated

and AM6701 treated hNAAA identified a peptide with mass of

1079.5177 Da (T-10b peptide; CTSIVAQDSR), containing the

catalytic cysteine in the control (untreated) sample, while a peptide

Table 1. Potencies of hNAAA inhibitors.

Inhibitor IC50 [14C] PEA (nM) IC50 PAMCA (nM) kinact (min21) KI (nM) kinact/KI (M21s21)

AM6701 7.262.5 7.760.2 0.04260.007 1363 55000

N-Cbz-serine b-lactone 19006500 17006200 0.02360.005 13006400 290

AM9023 6006130 350670 NA NA NA

The kinact and KI values for the covalent inhibitors were obtained as described in the Experimental Procedures. The IC50 values were calculated after 2 hourspreincubation of the enzyme and inhibitor before addition of the substrate. Values are averages 6 SD of three independent experiments.doi:10.1371/journal.pone.0043877.t001

Figure 2. Putative mechanism of inhibition of hNAAA for three compounds studied. Panel (A). Reversible inhibition of hNAAA by AM9023.Panel (B). Irreversible inhibition of hNAAA by AM6701 via thiocarbamylation of Cys126. Panel (C). Irreversible inhibition of hNAAA by N-Cbz-serine b-lactone most likely proceeds via route 2.doi:10.1371/journal.pone.0043877.g002



with a mass of 1150.5459 Da was observed only in inhibitor

treated samples (Figures 3a and 3b), with a concomitant decrease

in the intensity of the unmodified T-10b peptide peak. The

observed difference in mass, 71.0282 Da, between these two

peptides is equivalent to the mass of a dimethylcarbamyl group

(calculated 71.0371 Da) (Table 2). To confirm that the

1150.5459 Da ion was the T-10b peptide and determine which

amino acid was covalently modified with a dimethylcarbamyl

group, we performed an MS/MS analysis. The fragmentation

data confirmed that this was the T-10b peptide (Figure 4a), and

that the 71 Da additional mass was derived via carbamylation of

the N-terminal cysteine. The putative covalent mechanism of

AM6701 inhibition of hNAAA is shown in Figure 2b. To visualize

the enzyme active site modified by AM6701, we constructed a

hNAAA homology model based on the protein structure available

in the protein data bank that it has the greatest homology in

primary amino acid sequence with, which is with conjugated bile

acid hydrolase (CBAH) from Clostridium Perfringens. Only the b-

subunit was modeled with carbamylated catalytic nucleophile

Cys126 because it contains the putative catalytic triad, and the a-

subunit has no sequence homology with CBAH (Figure 5).

MALDI-TOF MS analysis of hNAAA inhibition by N-Cbz-serine b-lactone

Covalent modification by N-Cbz-serine b-lactone of the residue

His102, in addition to the catalytic Cys172 in the active site, of

hepatitis A virus 3C proteinase have been observed by NMR.

Alkylation of Cys172 or N-alkylation of His102 occurred in

hepatitis A virus 3C proteinase through the lactone cycle opening

at the b-carbon (Figure 2c, route 1) [35]. The most likely

mechanism of inhibition by N-Cbz-serine b-lactone with hNAAA

is that the catalytic Cys126 in hNAAA is alkylated (route 1) or

acylated (route 2) as shown in Figure 2c. A third possibility is

Cys126 carbamylation, resulting in only a fragment of the

compound covalently attached to the enzyme (Figure 2c, route

3). A slow regeneration activity of hNAAA inhibited with N-Cbz-

serine b-lactone in a rapid dilution assay suggested acylation or

carbamylation (route 2 and 3) rather than alkylation (route 1) of

Cys126 (Figure 2c). Analysis of the MALDI-TOF MS spectra for

the tryptic digests of untreated and N-Cbz-serine b-lactone treated

hNAAA identified a difference in mass of 221.0553 Da between

the peptides containing the catalytic cysteine in the control

(untreated) sample (1079.5177 Da; T-10b peptide; CTSI-

Figure 3. Tryptic digest fingerprint of purified hNAAA obtained by MALDI-TOF MS. Panel (A). Protein neat. Panel (B). AM6701 treatedenzyme. Panel (C). N-Cbz-serine b-lactone treated enzyme. The T10-b peptide (sequence: CTSIVAQDSR, theoretical mass 1079.515 Da) peakcontaining the catalytic nucleophile cysteine and its covalently modified forms are marked with an asterisk in each panel.doi:10.1371/journal.pone.0043877.g003

Figure 4. MALDI-TOF MS/MS analysis of the hNAAA tryptic peptide T10-b after covalent modification. Tandem MALDI-TOF MS/MSspectra of the T10-b peptide (sequence: CTSIVAQDSR) demonstrates covalent modification of Cys126 by both AM6701 (Panel (A)) and N-Cbz-serine b-lactone (Panel (B)).doi:10.1371/journal.pone.0043877.g004



VAQDSR) and inhibitor treated sample (1300.5730 Da), as

shown in Figures 3a and 3c. The 1300.5730 Da ion was not

observed in the untreated sample, and the unmodified T-10bpeptide peak is still present but at a reduced intensity. The

221.0553 Da difference between these two peptides eliminated the

possibility of route 3 in Figure 2c (calculated difference

131.0219 Da) and strongly suggested that the addition of N-Cbz-

serine b-lactone (calculated 221.06888 Da, Table 2) must proceed

by route 1 or 2 as shown in Figure 2c. The amino acid sequence of

1300.5730 Da ion identified by MS/MS analysis, as shown in

Figure 4b, was confirmed to correspond to the T-10b peptide with

Cys126 modified by the inhibitor. Our kinetic data demonstrating

the low in vitro stability of N-Cbz-serine b-lactone treated hNAAA

supports with the previous suggestion that a thioester bond is

formed after attack of sulfur at the 2-carbonyl [11], as this is a

more labile bond than the alkyl bond formed if the attack were at

the 4-methylene, and hence is strong evidence that inhibition

occurs by cysteine acylation via route 2 of Figure 2c. The

homology model of hNAAA with the N-Cbz-serine b-lactone

modified catalytic nucleophile Cys126, via acylation, is shown in

Figure 6.

In the course of preparing this manuscript it was reported by

Armirotti et al. that the b-lactones inhibit NAAA by S-acylation of

the catalytic N-terminal cysteine [36], confirming our data

presented in this manuscript and at the 2011 International

Cannabinoid Research Society meeting [37].

Conclusion

An understanding of structural organization and catalytic

mechanism of the human enzyme N-acylethanolamine-hydrolyz-

ing acid amidase is prerequisite to advance the development of

medicines with anti-inflammatory, analgesic and neuroprotective

properties. As the first step to hNAAA active site characterization

we applied an MS-based ligand-assisted protein structure

approach (LAPS) to identify an amino acid residue(s) in hNAAA

susceptible to selected irreversible inhibitors. To obtain a sufficient

amount of enzyme for the development, validation and executing

of HTS inhibitor assays we further optimized a previously

established HEK293-based hNAAA expression system to produce

three-fold more secreted functional protein. Different classes of

hNAAA inhibitors were pulled out during HTS screening of

compound libraries using a 3 point fluorescence based assay, and

the most potent were characterized further in a novel 8 point assay

for reversible (based on IC50 values) and irreversible (based on

kinact/KI values) hNAAA inhibitors. The mechanisms of hNAAA

inactivation by AM9023, AM6701 and N-Cbz-serine b-lactone

were investigated in biochemical and MS experiments. The

kinetics of hNAAA inhibition by AM9023 and MS analysis of

untreated and AM9023 treated hNAAA strongly suggest that this

isothiocyanate based compound is a reversible and non-covalent

inhibitor of hNAAA. AM6701 and N-Cbz-serine b-lactone inhibit

hNAAA in a covalent, time-dependent, and in the former case,

irreversible manner. We observed slow partial activity recovery of

hNAAA treated with N-Cbz-serine b-lactone, but not with

AM6701 in a rapid dilution assay. MS analysis of untreated and

Figure 5. Representation of the active site of hNAAA aftertreatment with AM6701. Homology model illustrates thiocarbamyla-tion of catalytic nucleophile Cys126 after treatment with AM6701.doi:10.1371/journal.pone.0043877.g005

Table 2. Mass of tryptic peptide containing Cys126 of hNAAA after covalent modification.

Peptide sequence Compound m/z calculated m/z measured Error (ppm)

CTSIVAQDSR DMSO (control) 1079.5150 1079.5177 2.5

CTSIVAQDSR AM6701 1150.5521 1150.5459 25.4

CTSIVAQDSR N-Cbz-serine b-lactone 1300.5838 1300.5730 28.3

T10-b peptides identified in the tryptic digest of untreated (control) and AM6701 or N-Cbz-serine b-lactone treated hNAAA samples.doi:10.1371/journal.pone.0043877.t002

Figure 6. Representation of the active site of hNAAA aftertreatment with N-Cbz-serine b-lactone. Homology model illus-trates acylated catalytic nucleophile Cys126 after treatment with N-Cbz-serine b-lactone.doi:10.1371/journal.pone.0043877.g006



AM6701 or N-Cbz-serine b-lactone inhibitor treated hNAAA

samples, following trypsin digestion, identified modification only

for the N-terminal cysteine (Cys126) of the b-subunit. These

experiments confirm that hNAAA belongs to the cysteine N-

terminal nucleophile class of enzymes, with Cys126 being the

critical residue in the active site susceptible to covalent inhibitors,

and establish methods to rapidly and efficiently determine the

covalent or reversible nature of NAAA inhibitors and determine

the potency of both types of inhibitors.

Author Contributions

Conceived and designed the experiments: JMW NZ SKV AM. Performed

the experiments: JMW NZ KMW SKV ALB. Analyzed the data: JMW NZ

AM. Contributed reagents/materials/analysis tools: JMW NZ KMW SKV

ALB AM. Wrote the paper: JMW NZ KMW SKV ALB AM.

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