Embed Size (px)
Citation preview
BIOCHEMICAL AND FUNCTIONAL STUDIES OF SYNAPTAMIDE
(N-DOCOSAHEXAENOYLETHANOLAMINE),
A METABOLITE OF DOCOSAHEXAENOIC ACID (DHA)
THESIS PRESENTED BY SHILPA SONTI
TO THE BOUVÉ GRADUATE SCHOOL OF HEALTH SCIENCES IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE
DEGREE OF DOCTOR OF PHILOSOPHY IN PHARMACEUTICAL SCIENCES WITH
SPECIALIZATION IN PHARMACOLOGY
NORTHEASTERN UNIVERSITY BOSTON, MASSACHUSETTS
December 13, 2016
2
Northeastern University
Bouvé College of Health Sciences
Dissertation Approval Dissertation title: BIOCHEMICAL AND FUNCTIONAL EFFECTS OF SYNAPTAMIDE,
(N-DOCOSAHEXAENOYLETHANOLAMINE), A METABOLITE OF
DOCOSAHEXAENOIC ACID (DHA) Author: SHILPA SONTI
Program: Doctor of Philosophy in Pharmaceutical Sciences with a specialization in
Pharmacology Approval for dissertation requirements for the Doctor of Philosophy in: Pharmaceutical Sciences
Dissertation Committee (Chairman) Date
Other committee members:
Date.
Date.
Date.
Date.
Date. __
Dean of the Bouve College Graduate School of Health Sciences:
Date.
Samuel Gatley
Ralph Loring
Robert Campbell
Barbara Shukitt-Hale
Jeanine Mount
Jonghan Kim
Ralph Loring
3
GLOSSARY
ABSTRACT 6
ACKNOWLEDGEMENTS 7
LIST OF ABBREVIATIONS 8
LIST OF TABLES 10
LIST OF FIGURES 11
A BACKGROUND AND SIGNIFICANCE 14-34
A.1 SIGNIFICANCE 14
A.2 INTRODUCTION TO LIPIDS 14
A.3 FATTY ACIDS 18
A.4 DOCOSAHEXAENOIC ACID 21
A.4.1 SYNTHESIS, UPTAKE AND RELEASE OF DHA 21
A.4.2 ROLE OF DHA IN MEMBRANE SYNTHESIS AND MODULATION 25
A.4.3 ROLE OF DHA IN NEURITOGENESIS AND SYNAPTOGENESIS 27
A.5 IS DHA RESPONSIBLE FOR ALL THE EFFECTS SEEN WITH ITS SUPPLEMENTATION?
28
A.6 SYNAPTAMIDE (N-DOCOSAHEXAENOYLETHANOLAMINE) 30
A.6.1 SYNTHESIS, UPTAKE AND METABOLISM 31
A.6.2 FUNCTIONAL ROLE OF SYNAPTAMIDE 32
B. RATIONALE 35
C MATERIALS AND GENERAL METHODS 38-41
C.1 ANIMALS 38
C.2 RADIOACTIVE COMPOUNDS 38
C.3 CHEMICALS 38
C.4 EQUIPMENT 39
C.5 N27 CELL CULTURE 39
C.6 STATISTICAL ANALYSIS 41
D SYNTHESIS OF SYNAPTAMIDE 42-57
D.1 INTRODUCTION 42
D.2 METHODS 45
D.2.1 SYNTHESIS OF N-ACYLETHANOLAMINES IN MOUSE BRAIN HOMOGENATES
45
D.2.2 SYNTHESIS OF N-ACYLETHANOLAMINES IN N27 CELLS 46
D.2.3 LIPID EXTRACTION 46
D.2.4 RADIO-TLC ANALYSIS 46
D.3 RESULTS 47
D.3.1 SYNAPTAMIDE IS SYNTHESIZED IN MOUSE BRAIN HOMOGENATES 47
D.3.2 IN VITRO SYNAPTAMIDE SYNTHESIS 49
D.4 DISCUSSION 54
D.5 CONCLUSION 56
E UPTAKE OF SYNAPTAMIDE– IN VITRO AND IN VIVO STUDIES 58-86
E.1 INTRODUCTION 58
E.2 METHOD 61
4
E.2.1 DETERMINATION OF TIME DEPENDENT SYNAPTAMIDE PARTITONING OVER TIME
62
E.2.2 ANALYSIS OF RADIOTRACER UPTAKE IN VIVO: MICRODISSECTION STUDIES
63
E.2.3 DETERMINATION OF RADIOTRACER UPTAKE IN VITRO 63
E.2.4 LIPID EXTRACTION FROM N27 CELLS 64
E.3 RESULTS 65
E.3.1 IN VIVO SYNAPTAMIDE UPTAKE 65
E.3.1.1 EXOGENOUS SYNAPTAMIDE ENTERS THE BRAIN 65
E.3.1.2 EXOGENOUS SYNAPTAMIDE IS TAKEN UP DIFFERENTIALLY INTO DIFFERENT BRAIN REGIONS
67
E.3.1.3 [14
C]SYNAPTAMIDE DISTRIBUTION PATTERN IN MOUSE BRAIN IS
DIFFERENT FROM THAT OF [14
C]ANANDAMIDE 68
E.3.1.4 [14
C]SYNAPTAMIDE UPTAKE IN VIVO IS HIGHER THAN THAT OF
[14
C]DHA. 70
E.3.2 IN VITRO UPTAKE STUDIES 72
E.3.2.1 ANANDAMIDE AND SYNAPTAMIDE HAVE SIMILAR UPTAKE PROFILES IN N27 CELLS
72
E.3.2.2 ANANDAMIDE AND SYNAPTAMIDE UPTAKE IN UNDIFFERENTIATED CELLS IS REGULATED BY THEIR HYDROLYSIS.
75
E.3.2.3 ANANDAMIDE AND SYNAPTAMIDE UPTAKE INTO UNDIFFERENTIATED N27 CELLS IS SIMILAR TO THAT OF AA AND DHA BUT NOT IN DIFFERENTIATING CELLS.
79
E.4 DISCUSSION 81
E.5 CONCLUSION 86
F ROLE OF FAAH ON SYNAPTAMIDE METABOLISM 87-111
F.1 INTRODUCTION 87
F.2 METHOD 88
F.2.1 MICRODISSECTION STUDIES WITH FAAH 88
F.2.2 FAAH ACTIVITY ASSAY 89
F.2.3 COMPETITION BINDING ASSAY 90
F.2.4 LIPID EXTRACTION 91
F.2.4.1 LIPID EXTRACTION FROM BRAIN HOMOGENATES 91
F.2.4.2 LIPID EXTRACTION FROM N27 CELLS 91
F.2.5 RADIO-TLC ANALYSIS 92
F.2.5.1 ONE-DIMENSIONAL TLC 92
F.2.5.2 TWO-DIMENSIONAL TLC 92
F.3 RESULTS 93
F.3.1 ROLE OF FAAH ON SYNAPTAMIDE UPTAKE IN VIVO 93
F.3.1.1 EFFECT OF FAAH INHIBITOR ON CRAIN AND BLOOD CARBON-14
LEVELS AFTER [14
C]SYNAPTAMIDE 93
F.3.2 ROLE OF FAAH ON SYNAPTAMIDE UPTAKE IN VITRO 94
F.3.2.1 SYNAPTAMIDE UNDERGOES HYDROLYSIS BY FAAH 94
F.3.2.2 THE HYDROLYSIS OF SYNAPTAMIDE BY FAAH IS SPONTANEOUS AS 97
5
WELL AS TISSUE MEDIATED.
F.3.2.3 SYNAPTAMIDE INHIBITS ANANDAMIDE HYDROLYSIS IN CRUDE BRAIN HOMOGENATES.
98
F.3.3 END FATE OF SYNAPTAMIDE 100
F.3.3.1 SYNAPTAMIDE INCORPORATES INTO PHOSPHOLIPIDS IN VIVO. 100
F.3.3.2 SYNAPTAMIDE PARTITIONS INTO PHOSPHOLIPIDS IN VITRO. 101
F.3.3.3 PHOSPHOLIPID PARTITIONING OF [14C-ETHANOLAMINE]SYNAPTAMIDE VERSUS [14C-DOCOSAHEXAENOYL]SYNAPTAMIDE
105
F.4 DISCUSSION 107
F.5 CONCLUSION 110
G FUNCTIONAL EFFECT OF EXOGENOUS DHA AND SYNAPTAMIDE ON NEURITOGENESIS
112-128
G.1 INTRODUCTION 112
G.2 METHOD 114
G.2.1 NEURITE ANALYSIS 115
G.3 RESULTS 115
G.3.1 EFFECT OF SYNAPTAMIDE AND DHA ON TOTAL NEURITE LENGTH IN DIFFERENTIATED N27 CELLS.
117
G.3.2 EFFECT OF SYNAPTAMIDE ON INDIVIDUAL NEURITE LENGTH OF DIFFERENTIATED N27 CELLS
120
G.3.3 EFFECT OF SYNAPTAMIDE ON THE NUMCER OF INDIVIDUAL NEURITES IN DIFFERENTIATED N27 CELLS
122
G.3.4 SYNAPTAMIDE AND DHA UPTAKE AND METABOLISM MAY CONTRIBUTE TO THEIR EFFECT ON NEURITOGENESIS AND NEURITE ELONGATION
123
G.4 DISCUSSION 125
G.5 CONCLUSION 127
H CONCLUDING REMARKS 129-132
I REFERENCES 133-150
6
ABSTRACT
The metabolite of docosahexaenoic acid (DHA), synaptamide, is reported to mediate the role of
DHA in neuritogenesis and synaptogenesis. Our long term goal is to identify the mechanism of
action involved in the purported neurogenic potential of synaptamide. In order to shed light on
its mechanism, gaps existing in the knowledge of synaptamide biochemistry have to close.
Thus, the objective of this dissertation is to investigate the fate of synaptamide in brain using
both in vivo and in vitro approaches.
The rationale behind this research is that, understanding the fate of synaptamide may aid in the
development of pharmacological interventions which might prove to be effective means in
overcoming neurological deficits. The specific aims for the proposed research are: 1) to
investigate the biosynthesis of synaptamide in vivo and in vitro; 2) To examine in vivo and in
vitro uptake of synaptamide; 3) To determine the role of FAAH in synaptamide metabolism and
its partitioning into phospholipids; 4) To analyze and compare a functional effect of synaptamide
with that of DHA.
This research is significant because resolving the metabolic fate of synaptamide can facilitate
the development of testable hypotheses which might eventually lead to the elucidation of its
biochemistry and signaling functions.
7
ACKNOWLEDGEMENTS
First of all, I would like to thank my advisor Dr. Gatley for supporting me and guiding me during
my PhD study. He was an incredible mentor to me constantly motivating me and guiding me
through this journey. He was an inspiration to us all and encouraged me to come up with my
own ideas and never discouraged me from shooting at the stars. Without him, this project might
have never taken shape. He made spending a substantial six years at Northeastern University a
memorable journey.
I would also like to thank my committee members: Dr. Loring, Dr. Kim, Dr. Shukitt-Hale and Dr.
Campbell. Their input and suggestions during my proposal and progress report meetings
contributed significantly in shaping up my dissertation. In addition, I would also like to thank all
the past and present members of our lab who helped me during various stages of my project. I
would particularly like to thank Kun Hu who had always been my sounding board to come up
explanations and new experiments and Mansi Tolia who helped me immensely during the final
stages of my work. I would also like to thank the Office of Science (CER), U.S. Department of
Energy who partly supported my research.
None of this would have been possible without the support of my family. My parents‘ hard work
and perseverance is the reason I was able to come to the United States and pursue my dream.
They empathized with me during my every success and failure and constantly motivated me to
think positively. Their words of encouragement always gave me the stimulation I needed during
the long hours spent in lab. My younger brother, Siddharth has always been there for me,
offering me support whenever I experienced the ―PhD blues‖. I am extremely fortunate to have
my best friend as my husband! His patience, love and support have helped me keep my calm at
my most difficult times. He never let me feel I was alone in this journey, and for that I am ever so
grateful. Even my in-laws, who‘ve known me only for a short period, have been extremely
understanding of my ordeals. I would also like to make a special mention to my extended family
who always wished the very best for me. I honestly could not have asked for a better family!
Last but not least, I would like to thank all my friends here, as well as from India; especially
Namrata and Prisca, for truly being there for me when I needed them the most.
Finally, I would like to acknowledge my late uncle and Godfather: Dr. S.S.R. Murthy. He was my
inspiration and motivation behind pursuing a career in Science.
I dedicate this dissertation to my family.
8
LIST OF ABBREVIATIONS
%IA/g: percentage of injected activity per gram [14C-arachidonoyl]anandamide, [3H- arachidonoyl]anandamide: anandamide labeled at the arachidonoyl moiety [14C-EA]anandamide: anandamide labeled at the ethanolamine moiety [14C-ethanolamine]synaptamide: synaptamide labeled at the ethanolamine moiety [14C-docosahexaenoyl]synaptamide: synaptamide labeled at the docosahexaenoyl moiety 1D-TLC: One Dimensional Thin Layer Chromatography 2D-TLC: Two Dimensional Thin Layer Chromatography AA: arachidonic acid AA-CoA: arachidonoyl-coenzyme A ACh4: α/β-hydrolase 4 ACHD6, ACHD12: αβ-hydrolase domain containing protein - 6 and - 12 ACSL6/ ACSL4: acyl-CoA synthase 6 acyl-CoA synthase 4 Acyl-CoA: acyl-coenzyme A AEA: anandamide Akt: Protein kinase C ALA: Alpha linolenic acid ATP: Adenosine triphosphate BBB: Blood Brain Barrier CB1 receptor: cannabinoid receptor 1 CB2 receptor: cannabinoid receptor 2 CD36KO: cluster of differentiation 36 knock out CoA: Co-enzyme A COX: Cyclooxygenase COX-2: cyclooxygenase-2 cPLA2: Cytosolic Phospholipase A2 CPM: counts per minute CREC: cyclic AMP response element binding protein BYP450: Cytochrome P450 dbcAMP: Dibutyryladenosine 3′,5′-cyclic monophosphate sodium DC: differentiating N27cells DHA: docosahexaenoic acid DHEA: docosahexaenoylethanolamide DLU/mm2: digital light units per mm2 DMSO: dimethyl sulfoxide EA: ethanolamine EC50: concentration of a drug that gives half-maximal response ER: endoplasmic reticulum FAAH: Fatty Acid Amide Hydrolase FABP: fatty acid binding protein FAT/ CD36: fatty acid translocase in combination with cluster of differentiation 36 FCS: Fetal Bovine Serum FLAT: FAAH-like-anandamide-transporter GAP-43: Growth Associated Protein-43 GP-NAE: glycerophospho-N-acylethanolamine
9
GSK3C: glycogen synthase kinase 3C I.P.: intraperitoneal I.V.: intravenous IC50: concentration of an inhibitor where the response (or binding) is reduced by half IL-6: Interleukin 6 iPLA2: Inducible Phospholipase A2 LA: linolenic acid LOX: lipoxygenase LysoPC: lysophosphatidylcholine LYSO-PE: lyso-phosphatidylethanolamine MCP-1: Monocyte chemoattractant protein-1 Mfsd2a: Major Facilitator Superfamily Domain-containing protein 2A mTOR: mammalian target of rapamycin N27 cells: 1RBN27 cells NaCl: sodium chloride NAEs: N-acylethanolamines NAPE: N-acyl phosphatidylethanolamine NAPE-PLD: N-acyl phosphatidylethanolamine-selective phospholipase D NAT: N-acyltransferase N-DHPE: N-docosahexaenoylphosphatidylethanolamine NE-DHA: non-esterified DHA NGF: Nerve Growth Factor NPD1: neuroprotectin D1 PBS: Phosphate Buffered saline PC: phosphatidylcholine PE: phosphatidylethanolamine PET: positron emission tomography PF3845: FAAH inhibitor PI: Phosphatidylinositol PIP3: Phosphatidylinositol (3,4,5)-trisphosphate PLA/AT: phospholipase activity with O-acyltransferase activity PLA1: phospholipase A1 PLA2: phospholipase A2 PLC: phospholipase C PLD: phospholipase D PPAR-α: peroxisome proliferator-activated receptor alpha PS: phosphatidylserine PSD-45: postsynaptic density protein 45 PSS: phosphatidylserine synthase PUFA: polyunsaturated fatty acid RAR: retinoic acid receptor Rf: retention factor RXR: Retinoid X receptors SD: Standard Deviation
sn‑1 and sn‑2: nucleophilic substitution at position 1 or 2 of phospholipid
TLC: Thin-layer chromatography TME: Tris Magnesium EDTA UDC: undifferentiated N27 cells
10
LIST OF TABLES Table A.1: Various classes of lipids and some of their subclasses
Table D.1: Average % IA/g in blood, brain and urine of mice over one hour treatment
Table D.2: Absolute signal intensity ratios of radiotracer accumulation in some brain regions.
Table D.3: Ratios of % [14C]DHA uptake and % [14C]synaptamide uptake in various brain
regions
11
LIST OF FIGURES
Fig A.1: Classification of lipids based on their chemical structures
Fig A.2: Structure of an omega-3 fatty acid, Linolenic acid (18:3n-3)
Fig A.3: Biosynthesis of long chain polyunsaturated fatty acids.
Fig A.4: (A) Proposed schematic for uptake and incorporation of DHA into phospholipids. (C) A
schematic representing the transport of lysoPC-DHA across the BBB by Mfsd2a (Major
Facilitator Superfamily Domain-containing protein 2A) transporter
Fig A.5: Site of action of various phospholipases and the products formed.
Fig A.6: The role of DHA in the synthesis of phospholipid, phosphatidylserine and in membrane
modulation
Fig A.7: Summary of various metabolites of AA and DHA (and their functional implications)
released from membrane phospholipids by the action of PLA2
Fig A.8: Structure of synaptamide (N-docosahexaenoylethanolamine)
Fig D.1: Representative TLC used for the quantification of NAE synthesis.
Fig D.2: Quantification of TLCs of brain lipid extracts to determine NAE synthesis
Fig D.3: % radioactivity in chloroform extracts of N27 cells as determined from scintillation
counts
Fig D.4: Representative TLC images NAE synthesis from radiolabeled free fatty acid.
Fig D.5: (A and C) TLC Quantification of chloroform extracts of cells incubated with radiolabeled
fatty acids. (C) Anandamide synthesis in N27 cells from 200 nM [14C]arachidonic acid.
Fig D.6: Autoradiograph and charred TLC image representative of synaptamide synthesis in
N27 cells incubated with 200 nM [14C]DHA
Fig E.1: (A) Blood to brain ratio of [14C]synaptamide. (C) Average %IA/g radioactivity in Blood
and brain over time
Fig E.2: Analysis of the regional distribution of radiolabel in animals injected with 0.1μCi
[14C]synaptamide, 0.1μCi [14C]DHA and 1μCi [14C]ethanolamine (I.V.).
Fig E.3: The pattern of distribution of ethanolamine as a control as evaluated by
autoradiography
12
Fig E.4: Comparison of uptake versus hydrolysis with [14C]anandamide and [14C]synaptamide in
N27 cells.
Fig E.5: Comparison of uptake and hydrolysis with [14C]anandamide versus [14C]synaptamide in
N27 cells.
Fig E.6: Comparison of [14C]anandamide uptake and hydrolysis in N27 cells with or without
PF3845
Fig E.7: Comparison of [14C]synaptamide uptake and hydrolysis in N27 cells with or without
PF3845
Fig E.8: Comparison of [14C]arachidonic acid and [14C]DHA uptake in N27 cells
Fig E.9: Comparison of NAE uptake versus their corresponding fatty acid uptake in N27 cells
Fig F.1: brain regional concentrations of 14C in animals euthanized 15 min after injection of
0.1μCi [14C] synaptamide (i.v.), with or without PF3458 pretreatment
Fig F.2: Time dependent increase in radioactive counts in the aqueous layers of brain
homogenates treated with [14C]anandamide and [14C]synaptamide
Fig F.3: Time dependent increase in the hydrolysis of [14C]anandamide and [14C]synaptamide in
brain homogenates
Fig F.4: Time dependent hydrolysis of [14C]anandamide and [14C]synaptamide in brain
homogenates treated with or without PF3845
Fig F.5: Comparison of N-acylethanolamine hydrolysis in tissue mediated and tissue
independent environments
Fig F.6: Representative graph showing displacement of [14C] anandamide binding to FAAH in
mouse brain homogenates by unlabeled anandamide (AEA), synaptamide (DEA) or DHA plus
ethanolamine (EP).
Fig F.7: Representative 1D-TLC autoradiograph of brain (A) and Blood (C) lipid extract from
mice euthanized after 0, 5, 15, 30 and 60 minutes after the administration of exogenous
[14C]synaptamide.
Fig F.8: Representative TLC images of anandamide and synaptamide uptake.
Fig F.9: 2D-TLC of standard non-radioactive phospholipids, PS: phosphatidylserine (A) and PC:
phosphatidylcholine (C) and cell lipid extract of N27 cells incubated with [14C-EA] Synaptamide
(C).
13
Fig F.10: Quantification of TLC spots shows the partitioning of anandamide and synaptamide
into phospholipids of cells in the absence (A) or presence (C) of PF3845.
Fig F.11: Quantification of TLC spots shows the partitioning of [14C-EA] Synaptamide and [14C-
DHA] Synaptamide into phospholipids of cells in the absence (A) or presence (C) of PF3845
Fig G.1: Representative images of undifferentiated (A) and differentiated (C) N27 cells.
Fig G.2: The representative images and traces of differentiated N27 cells
Fig G.3: Estimation of total neurite length with increasing doses of either DHA or synaptamide
Fig G.4: Comparison of the Total neurite length of N27 cells treated with either DHA or
synaptamide.
Fig G.5: Representative graph of frequency distribution of total neurite length after either DHA
(A) or synaptamide (C) supplementation in differentiated N27 cells.
Fig G.6: Estimation of total length of individual neurites selected from randomly selected cells
supplemented with increasing doses of either DHA or synaptamide
Fig G.7: Representative graph of frequency distribution of individual neurite length after either
DHA (A) or synaptamide (C) supplementation in differentiated N27 cells
Fig G.8: Total number of neurites (sum of primary, secondary and tertiary Branches) from N27
cells (n=70) supplemented with either DHA or synaptamide
Fig G.9: Comparison of neurite lengths (A) and the number of neurites (C) from randomly
selected N27 cells (n=70) treated with either DHA 100 nM or synaptamide 100 nM.
Fig G.10: Time dependent phospholipid incorporation of exogenous [14C]DHA and [14C-
docosahexaenoyl]synaptamide in undifferentiated (A) and differentiating (C) N27 cells
14
A. BACKGROUND AND SIGNIFICANCE
A.1. Significance
Docosahexaenoic acid (DHA; 22:6n-3) and arachidonic acid (AA; 20:4) are the major fatty acids
of the brain with crucial roles in neurodevelopment and proper brain functioning. While some of
their downstream effects can be explained by a direct role, others may be explained by their
bioactive metabolites. Consistent with this, many metabolites of arachidonic acid and DHA have
neuroprotective functions (e.g. – neuroprotectin D1, NPD1). Synaptamide, a recently discovered
endocannabinoid-like molecule that incorporates DHA is reported to mediate the neuritogenic
effects of DHA. The mechanism mediating this effect is poorly understood. This lack of
knowledge hinders the development of pharmacological interventions which may prove to be
effective means in overcoming neurological deficits. In order to be able to elucidate its
biochemical pathways, it is first important to understand the fate of synaptamide in the neural
cell. As an outcome of the proposed investigations, we expect to have determined not only the
metabolic profile of synaptamide, but also the possibility that DHA (released from synaptamide)
mediates some of its effects. The proposed research is significant because once the metabolic
fate of synaptamide in brain is known, it will facilitate the development of testable hypotheses,
finally leading to the elucidation of its biochemistry and signaling functions.
A.2. Introduction to lipids
Lipids can be defined as hydrophobic constituents of the cell which comprise a large number of
molecules with various combinations of fatty acids conjugated with different backbone
15
structures. (Wenk, 2005). There are about 2000 different lipid species in mammalian cells, often
categorized into about 8 classes with a number of subclasses. The major categories and some
of their subclasses are listed in Table A.1 (Adibhatla et al., 2007).
Lipid class Subtypes
Fatty acyls Free fatty acids and conjugates; Eicosanoids; Docosanoids; Fatty
Alcohols, Aldehydes and Esters
Glycerolipids Mono-, di-, and triacylglycerol
Glycerophospholipids Phosphatidylcholine, Phosphatidylethanolamine, Phosphatidylserine,
Phosphatidylinositol
Sphingolipids beramide, Sphingomyelin, Glycosphingolipids (Gangliosides)
Sterol lipids Sterols including cholesterol, steroids, bile acids
Prenol lipids Isoprenoids, Polyprenols, Quinones
Saccharolipids Acylaminosugars, Acylaminosugar glycans
Polyketides Macrolide and Aromatic polyketides
Table A.1: Various classes of lipids and some of their subclasses.
Future Lipidol. 2007 Aug; 2(4): 403–422 (Adibhatla et al., 2007).
Lipids have important roles in normal neuronal biochemistry and physiology. They are integral
components of the plasma membrane that functions as a carrier between the intra and
extracellular compartments. Being inherent components of the membrane, they have a crucial
role in determining the localization of membrane proteins as well as in regulating their actions.
Lipids also influence important cellular functions such as exo- and endocytosis, information
16
relay within and beyond the cells, etc., (Muller et al., 2015). The major components of the
mammalian membrane lipid fraction are glycerophospholipids, sphingolipids and the sterol lipid
cholesterol; chemical composition of each lipid class varying greatly with the cell type and
membrane (van Meer et al., 2008) (fig A.1). These lipids, alone or in combination with other
lipids, have the ability to stabilize in different phases by interacting with the changing
environment and to adapt by modifying their chemical structures, by altering their fatty acid
composition and phospholipid head-groups. This ability enables lipids to modulate cellular
communication (Piomelli et al., 2007).
Fig A.1: Classification of lipids based on their chemical structures. Highlighted in red are the core-
structures from which the various lipid classes take their name. Highlighted in blue are the
functional groups from which the various lipid subclasses take their name. Abbreviations: PC,
17
phosphatidylcholine; PE, phosphatidylethanolamine; PS, phosphatidylserine; PG,
phosphatidylglycerol; PI, phosphatidylinositol; ber, ceramide; SM, sphingomyelin; Hexber,
hexosyl ceramide; Cho, cholesterol. (Paglia et al., 2015) open access
Glycerophospholipids have a glycerol backbone with fatty acids at sn-1 and sn-2 positions and a
head group at sn-3 position. An unsaturated fatty acid is usually conjugated at the sn-2 position
while a saturated fatty acid occupies the sn-1 position. Depending on the type of head group
occupying the sn-3 position, glycerophospholipids are categorized into phosphatidylcholines
(PC; glycerophosphocholines), phosphatidylethanolamines (PE;
glycerophosphoethanolamines), phosphatidylserines (PS; glycerophosphoserines) and
phosphatidylinositols (PI; glycerophosphoinositols). (Muller et al., 2015). Sphingolipids
constitute another major class of membrane lipids with a ceramide backbone. beramide consists
of a sphingosine molecule conjugated with a long chain saturated fatty acid. Sphingomyelin and
gangliosides are major sphingolipids in the brain (van Meer et al., 2002). Sterol lipids are non-
polar structural lipids and they integrate with glycerophospholipids and sphingolipids to form an
intricate cell membrane structure. The distribution of these 3 lipid classes varies with different
cell types and different organelles (van Meer et al., 2008).
A major factor that modifies membrane fluidity in the brain is the nature of the phospholipid
hydrophobic tail – the fatty acid residues linked to the sn‑1 and sn‑2 hydroxyl groups on the
glycerol backbone. At physiological temperatures, the length of a phospholipid molecule is
directly proportional to the number of carbon atoms and inversely proportional to the number of
double bonds present in its fatty acid chains (Piomelli et al., 2007).
18
A.3. FATTY ACIDS
Long-chain fatty acids are a subset of lipids, comprising of hydrocarbons chains of various
lengths and terminate with a carboxylic acid group. Their synthesis is initiated by the
condensation of malonyl coenzyme A units by fatty acid synthetase complex. In animals, long
chain fatty acid chain length varies between 14-22 carbons (Christie). They are subcategorized into
saturated, mono- and polyunsaturated fatty acids. Different classes of major structural lipids
differ among themselves based on their fatty acid composition eg., sphingolipids incorporate
very long chain saturated and mono- or di-unsaturated fatty acids, ceramides incorporate
monounsaturated fatty acids and phospholipids have both saturated and polyunsaturated fatty
acids (Christie). The unsaturated fatty acid component, conjugated on the sn-2 position, is an
important determinant of the molecular flexibility and other properties of phospholipids.
The varying degrees of unsaturation offered by different fatty acids allows cells to modulate
membrane structure and function. Biological fatty acids can be of various lengths, and the
naming convention is to indicate the position of double bonds with reference to the terminal
methyl group. Regardless of the chain length, this is labeled omega (―ω‖ or sometimes ―n‖).
Long chain polyunsaturated fatty acids are of three types – omega-3, omega-6 and omega-9.
The term ω-3 or n-3 means that the third carbon-carbon bond from the end of the chain (the
omega carbon) is a double bond chain (Fig A.2).
19
Fig A.2: Structure of an omega-3 fatty acid, Linolenic acid (18:3n-3)
(Accessed from http://psychology.wikia.com/wiki/Omega-3_fatty_acid on 11/15/2016)
The omega-3 and omega-6 fatty acids are considered as Essential Fatty Acids (EFAs) as their
biosynthesis requires precursors – linoleic acid (LA; 18:2n-6), linolenic acid (ALA; 18:3n-3) that
cannot be synthesized by animals and can therefore only be obtained through diet. Linoleic acid
and linolenic acid are synthesized in plants from stearic acid (SA; 18:0). Stearic acid is
converted to oleic acid (18:1 n-9) by Δ9- desaturase which is exported into the endoplasmic
reticulum where it is further acted upon by Δ12- and Δ15- desaturases to synthesize linoleic acid
and linolenic acid respectively (Huerlimann et al., 2014) (fig A.3). Higher mammals do not have
the desaturase enzymes (Δ9-, Δ12- and Δ15- desaturases) responsible for the conversion of oleic
acid into linoleic acid and linolenic acid (Pereira et al., 2003). This is the reason why linoleic acid
and linolenic acid are considered essential and have to be procured through diet.
20
Fig A.3: Biosynthesis of long chain polyunsaturated fatty acids.
Gene, Volume 545, Issue 1, 2014, 36–44 (Huerlimann et al., 2014) with permission
Fatty acid desaturase enzymes are capable of introducing unsaturation in specific positions of
the fatty acid chains – from either the (tail) methyl- end of the chain or the carboxylic head
group-(front) end of the chain (Park et al., 2009). Cloth kinds of desaturase enzymes along with
elongases synthesize longer chain fatty acids (arachidonic acid (AA; 20:4n-6) and
docosahexaenoic acid (DHA; 22:6n-3)) from precursor fatty acids. Plants and lower animals
such as birds have both kinds of desaturase enzymes while humans only express ―front end‖
enzymes (Nakamura et al., 2004),(Tocher et al., 1998).
21
A.4. Docosahexaenoic acid (DHA; 22:6n-3)
A.4.1. Synthesis, uptake and release of DHA
Essential fatty acids make up about 20% of the dry weight of the brain. One third of these
belong to the omega-3 PUFA family. DHA is the most abundant omega-3 polyunsaturated fatty
acid in the brain (Contreras et al., 2000). DHA accumulation in brain is gradual starting from the
third trimester of pregnancy until the brain is completely developed (Clandinin et al., 1980). It
was shown that brain ω3: ω6 ratio (contributed mainly by DHA) in the phospholipid
(phosphatidylethanolamine) component increased with age in children (children > toddler >
infant > fetus) (Martinez et al., 1998). This age-dependent DHA accretion pattern during brain
development was consistent over a wide variety of species including rats (Green et al., 1996)
and piglets (Purvis et al., 1982).
Biosynthesis of DHA takes place mainly in liver endoplasmic reticulum from linolenic acid
through chain elongation and desaturation (Sprecher, 2000) (fig 2). Δ4, Δ5 and Δ6-desaturases
are most important for this conversion (Kim, 2007). As the essential fatty acid precursors are
actively converted to DHA; its levels are never completely depleted. Amongst brain cells
(neurons, astrocytes, microglia, and oligodendrocytes), only astrocytes can synthesize DHA
(Moore et al., 1991). Neurons lack the Δ4 desaturase enzyme essential for DHA synthesis.
Astroglial DHA synthesis is negatively influenced by the availability of preformed DHA (Williard
et al., 2001) and thus may represent a quantitatively minor source for the neural DHA accretion,
making its procurement from diet very important, especially during embryonic development.
22
It was assumed until recently that transport of DHA to the brain occurs from the free non-
esterified DHA (NE-DHA) bound to serum albumin. The NE-DHA would then cross the BBB as a
result of competition between the hydrophobic domain of albumin and that of the endothelial
layer (Picq et al., 2010). However, a recent article reports that DHA (and other polyunsaturated
fatty acids) is more efficiently taken up by the developing brain when esterified in
lysophosphatidylcholine (lysoPC-DHA) as compared to the non-esterified form, both forms
being bound to albumin (Lagarde M, 2000). In contrast, albumin bound NE-DHA was
preferentially taken up by heart and liver (Fig A.4A). Lysophosphatidylcholine, the major lipid
component of plasma but a minor component of phospholipids in cells and tissues, is derived
from phosphatidylcholine (PC) and is amphiphilic (Croset, 2000). It is expected that DHA will be
esterified at the sn-2 position of the lysophosphatidylcholine because the phosphatidylcholine
precursors have polyunsaturated fatty acids conjugated at this position. However, it has been
shown that sn-1-lyso-sn-2-DHA-phosphatidylcholine isomerizes at neutral pH to the more
thermodynamically stable sn-1-DHA-sn-2-lysophosphatidylcholine, and the ratio of the two
isomers depends on their half lives in plasma and on catabolism in tissues (Croset, 2000). While
both isomers are taken up by the brain in vivo to the same extent (Morash S C, 1989), the
mechanism by which they can cross the BBB was unknown until recently (Fig A.4A). The
metabolic fates of the isomers differ and also depend on the cell type. Both isomers are
reacylated in glial cells whereas sn-1-DHA-sn-2-lysophosphatidylcholine is deacylated and sn-
1-lyso-sn-2-DHA-phosphatidylcholine is reacylated in neurons (Morash S C, 1989). The
reacylation of either isoform forms sn-1-DHA-sn-2-DHA-phosphatidylcholine which is a
substrate for acyltransferases and phospholipases. Deacylation of either isomer forms
glycerophosphocholine, a source for choline in the brain.
23
A B
FigA.4: (A) Proposed schematic for uptake and incorporation of DHA into phospholipids. DHA
produced from linolenic acid (LNA) in the liver is incorporated in lipoproteins, phospholipids and
triglycerides. Enzymes including phospholipase A1, endothelial lipase and lipoprotein lipase
generate lysoPC-DHA which is then taken up by the brain to be re-acylated and/or hydrolyzed to
provide DHA for incorporation into brain phospholipids. Interconversion between brain
phospholipids may also occur (Picq et al., 2010) reused with permission. (B) A schematic
representing the transport of lysoPC-DHA across the BBB by Mfsd2a (Major Facilitator
Superfamily Domain-containing protein 2A) transporter. (LysoPC-DHA can either be sn-2-DHA-sn-
1-lysophosphatidylcholine or sn-1-DHA-sn-2-lysophosphatidylcholine).
Possible mechanisms proposed for transport of fatty acids across the BBB to the brain are
diffusion and facilitated transport (Qi et al., 2002). Fatty acids may diffuse across membranes
using the ―flip flop‖ mechanism. This was observed in transport of fatty acids across the cell
membrane and other unilamellar membranes, but ―flip flop‖ transport across the BBB is poorly
characterized (Hamilton et al., 2001). Additionally, membrane proteins such as fatty acid
translocase in combination with cluster of differentiation 36 (FAT/ CD36) and cytosolic proteins
24
such as fatty acid binding proteins (FABPs) are shown to facilitate the transport of fatty acids
(Abumrad et al., 1999). While CD36 dependent FATP was found to be most effective in fatty
acid transport; transport of DHA remained unchanged in CD36KO animals (Lo Van et al., 2016)
suggesting that other transporters maybe involved in DHA transport. It has been recently shown
that Mfsd2a (Major Facilitator Superfamily Domain-containing protein 2A), a protein expressed
in BBB endothelium, is responsible for the selective transport of DHA incorporated in
lysophosphatidylcholine over other forms of DHA or NE-DHA (Nguyen et al., 2014). Fig A.4C
demonstrates the possibility of selective transport of DHA incorporated in
lysophosphatidylcholine (either isomer) bound to albumin across the BBB into the brain it can
undergo metabolism.
Fig A.5: Site of action of various phospholipases and the products formed. Phospholipase A1 and
A2 cleave at sn-1 and sn-2 positions respectively to produce the corresponding free fatty acids
and lysophospholipids. Phospholipase C cleaves the phospholipid to form diacylglycerol and
Phospholipase D cleaves to form phosphatidic acid. R1 and R2 represent free fatty acids. X refers
to head groups (choline, ethanolamine, inositol, etc.) of the phospholipid. Accessed from
https://www.hindawi.com/journals/er/2011/392082/fig1/ on 11/15/16
25
Phospholipases release free fatty acids – Arachidonic acid and DHA from phospholipids.
Depending on the site of their action, they are categorized into four classes – phospholipase A1
(PLA1; acts at sn-1 position of phospholipid), phospholipase A2 (PLA2; acts at sn-2 position of
phospholipid), phospholipase C (PLC) and phospholipase D (PLD) (Fig A.5). Since
polyunsaturated fatty acids are preferentially esterified at the sn-2 position of the phospholipids,
phospholipase A2s are responsible for releasing the free fatty acid. Studies using radiolabelled
fatty acids show in vitro and in vivo selectivity of cPLA2 and iPLA2 to arachidonic acid and DHA
respectively (Rapoport et al., 2011). DHA released from the ER is activated by the action of
acyl-CoA synthetase and acyltransferase to form docosahexaenoyl-CoA which is then
incorporated stereospecifically at the sn−2 position of new membrane phospholipids. The
enzyme involved in DHA acylation is Acsl6 (Marszalek et al., 2005a), which exhibits a low
affinity for this substrate (Km = 26 μM) (Reddy et al., 1984) relative to usual brain DHA levels
(1.3–1.5 μM) (Contreras et al., 2000). Some DHA that escapes this pathway is subjected to a
number of catabolic pathways resulting in bioactive metabolites such as docosanoids, resolvins,
etc. (Rapoport et al., 2007).
A.4.2. Role of DHA in membrane synthesis and modulation
DHA and arachidonic acid are the major polyunsaturated fatty acids in the brain. The half-life of
DHA in Blood of healthy human subjects is 20 ± 5.2 hours (Pawlosky et al., 2001) and 22.4 ±
2.9 hours in brain phosphatidylcholine which is much longer than that of AA (3.79 ± 0.12 hours)
implicating a preferential incorporation of DHA into phosphatidylcholine (Rapoport, 2005). DHA-
phosphatidylcholine can undergo transesterification to produce DHA-phosphatidylethanolamine.
26
brain phosphatidylcholine or phosphatidylethanolamine can in turn produce phosphatidylserine
(PS) by Serine-Base-Exchange reaction mediated by PSS (phosphatidylserine synthase)
enzymes (Vance et al., 2004) (Fig A.6).
Fig A.6: The role of DHA in the synthesis of phospholipid, phosphatidylserine and in membrane
modulation (Adapted from Kim HY, OCL 2011; 18(5): 237-241) (Kim, 2007) reused with permission.
Phosphatidylserine represents the major negatively charged phospholipid class in mammalian
cell membranes, where it is localized mainly on the cytosolic side. Brain PSS enzymes
demonstrate a strong preference for DHA-containing phospholipids (18:0, 22:6n-3 > 18:0,
22:5n-6 > 18:0, 18:1 > 18:0, 20:4n-6 species) (Kim et al., 2004) for the synthesis of
phosphatidylserine. DHA enrichment in vitro can induce phosphatidylserine accumulation in
neuronal cells, primarily because of the accumulation of 18:0, 22:6-PS (Akbar et al., 2005).
27
Diacylglycerol species containing DHA or arachidonic acid are preferentially utilized for
phosphatide synthesis as opposed to triglyceride synthesis (Marszalek et al., 2005b). Hence,
dietary DHA would increase the accumulation of 18:0, 22:6- phosphatidylserine in the
membrane which is responsible for healthy physiological functions. Phosphatidylserine
participates in constitutive cell signaling by interacting with important signaling proteins for their
activation. For example, interaction between protein kinase C and Raf-1 is modulated by levels
of phosphatidylserine in the membrane. Phosphatidylserine favors Akt translocation and
promotes cell surviving signals such as PIP3, thus playing a role in neuronal survival (Akbar et
al., 2005) (Fig A.6). This is particularly significant in suboptimal conditions, where the generation
of survival signals (PIP3) is limited.
A.4.3. Role of DHA in neuritogenesis and synaptogenesis
Neurogenesis (the generation of neurons) involves both proliferation and differentiation.
Neuronal differentiation in turn involves neuritogenesis and synaptogenesis. ‗‗Neurites‘‘ are
precursors of axons and dendrites that, once formed, serve to polarize the neuron. A growth
cone is the mobile tip of the neurite specialized for elongation (Clagett-Dame et al., 2006). DHA
promotes neuronal differentiation by influencing neurite growth and synaptogenesis (Kan et al.,
2007). The production of phospholipids and other membrane components is one of the pre-
requisites for neurite growth (Banker, 1996). Omega-3 polyunsaturated fatty acids present in
growth cones and synaptosomal membranes (Youdim, 2000) enable them to play a significant
role in the dynamics of synapses. When supplemented with DHA, the DHA-favored
phospholipids are incorporated in neuronal membranes and thus can influence the quaternary
structure of membrane proteins. DHA supplementation increased the levels of an axonal growth
28
marker, growth associated protein-43 (GAP-43), a protein associated with growth cone
formation (Banker, 1996). DHA influences synapse formation through Retinoid X receptors
(RXRs). Retinoid X Receptor forms heterodimers with nuclear receptors such as retinoic acid
receptor (RAR), peroxisome proliferator-activated receptors (PPARs), or Nurr1 for
transcriptional regulation of target genes (Aarnisalo et al., 2002; Lane et al., 2005). These
dimers play an important role in neurodevelopment by regulating genes involved in the control
of synaptic plasticity, cytoskeleton, and membrane assembly, as well as signal transduction and
ion channel formation (Lane et al., 2005) (Maden, 2002). Being an endogenous ligand for
Retinoid X Receptor; DHA binds to Retinoid X Receptor within the functional dimer enhancing
its transcriptional activity. Nurr1–RXR signaling prevents the loss of synaptic proteins as DHA
supports neuronal survival, particularly of dopaminergic neurons in embryonic stage by binding
to RXR and influencing the formation of Nurr1–RXR heterodimers. These studies suggest
neuritogenic and synaptogenic roles for DHA.
A.5. Is DHA responsible for all the effects seen with its supplementation?
While there is evidence that indicates direct roles of DHA in bringing about some of its effects,
the possible involvement of its metabolites cannot be ignored. The free fatty acids released from
membrane phospholipids that escape acylation and re-incorporation can be subjected to
enzymatic or non-enzymatic catabolism. cyclooxygenase (COX), lipoxygenase (LOX),
cytochrome P450, and probably other enzymes can generate biologically active metabolites
from arachidonic acid and DHA that participate in signal transduction mediating some of their
beneficial effects (Phillis, Horrocks et al. 2006) (Fig A.7).
29
30
Fig A.7: Summary of various metabolites of AA and DHA (and their functional implications)
released from membrane phospholipids by the action of PLA2. (Adapted from Tai EKK, Food
Funct., (2013)4, 1767-1775) (Tai et al., 2013) reused with permission.
LOX metabolites generated from arachidonic acid, in particular, have pro-inflammatory effects
(Piomelli, 1994). LOX-metabolites of DHA (resolvins and protectins to name a few) also serve
as signaling molecules accounting for anti-inflammatory effects of DHA. Non-enzymatic free
radical-mediated peroxidation of free fatty acids generates prostaglandin-like compounds – F2-
isoprostanes from arachidonic acid and F4-neuroprostanes from DHA respectively (Montuschi
et al., 2004). The levels of F4-neuroprostane levels in human brain and cerebrospinal fluid are
elevated in Alzheimer‘s disease suggesting that peroxidation products of DHA may serve as
potential biomarkers (Montuschi et al., 2004). DHA is transformed by a 15-LOX-like enzyme to
(10,17S)-docosatriene (Hong et al., 2003), also termed neuroprotectin D1 (NPD1) because of its
neuroprotective properties. In an Alzheimer's disease model, NPD1 suppressed αβ-42-induced
neurotoxicity by inducing neuroprotective and anti-apoptotic gene expression (Lukiw et al.,
2005). Synaptamide, an endocannabinoid-like metabolite of DHA was recently proposed to play
a role in mediating the neuritogenic effects of DHA (Kim et al., 2011). While some effects of
DHA can be explained by the formation of oxidative metabolites, the notion that neuritogenic
effects of DHA are mediated by synaptamide can only be evaluated if the biosynthesis and
enzymatic hydrolysis of this biochemical are understood.
A.6. Synaptamide (N-docosahexaenoylethanolamine)
31
A.6.1. Synthesis, uptake and metabolism
Fig A.8: Structure of synaptamide (N-docosahexaenoylethanolamine)
Synaptamide (N-docosahexaenoylethanolamine) is found in brain tissue at levels comparable
with its structural analog, anandamide (N-arachidonoylethanolamine) (Bisogno T, 1999) (fig
A.8). The metabolic pathways involved in synaptamide biosynthesis remain unclear; however, it
has been proposed that it is biosynthesized from its corresponding N-acyl
phosphatidylethanolamine (NAPE) through a single NAPE-PLD-dependent pathway (NAPE-
PLD) (Schmid et al., 1996) similar to anandamide. The presence of didocosahexaenoyl
phosphatidylethanolamine or phosphatidylcholine, N-acyl transferase and N-DHPE (N-
docosahexaenoylphosphatidylethanolamine), the precursors for synaptamide synthesis have
been identified in the bovine retina (Bisogno T, 1999). The synthesis of synaptamide from DHA
was reported in hippocampal neurons (Cao et al., 2009), cortical neurons and neural stem cells
(Rashid et al., 2013); however, the mechanism of its synthesis and the presence of its
precursors in neural tissue is yet to be validated. It is not known whether synaptamide is
synthesized in the neuronal compartment or the glial compartment. Concurrently it is also
32
unknown whether synaptamide given exogenously can transfer into the intracellular
compartment of neuronal cells. Studies suggest that a high-affinity transport system exists to
transport anandamide to neurons (Hillard et al., 2003). Based on the structural similarity with
anandamide, it is reasonable to assume that neurons can take up synaptamide and that intact
synaptamide might be present inside the cell for a period of time prior to its hydrolysis. It has
been proposed that hydrolysis of synaptamide releases free DHA and ethanolamine (Bisogno T,
1999). Inhibition of FAAH (Fatty Acid Amide Hydrolase) purportedly increased the functional
effect of synaptamide in vitro (Kim et al., 2011). Based on these reports, it can be assumed (but
it has not been confirmed, to our knowledge) that synaptamide may be a substrate for the
enzyme FAAH that hydrolyses anandamide. Intact synaptamide may also undergo enzymatic
oxidation to form novel bioactive products (Yang et al., 2011) with hypothesized anti-
inflammatory and anti-apoptotic roles.
A.6.2. Functional role of synaptamide
Synaptamide inhibited forskolin-mediated cAMP production (IC50 = 6 μM) in CHO–HCR cells
(Felder et al., 1993). Lipopolysaccharide-induced IL-6 and MCP-1 production was suppressed
by synaptamide in 3T3-L1 pre-adipocytes, suggesting an anti-inflammatory role in adipose
tissue (Balvers et al., 2010). Synaptamide also decreased the viability of the LNCaP and PC3
prostate cancer cell lines (IC50 120–130 μM) (Brown et al., 2010). Synaptamide levels in E-18
fetal hippocampi (155 ± 35 fmol/μmol) are significantly higher than the anandamide level (44±3
fmol/ μmol). This relatively high content suggests that synaptamide might have an important role
in the hippocampus. In support of this observation, synaptamide was found to stimulate neurite
growth, synaptogenesis and synaptic protein (synapsin-1, synaptophysin, PSD-45) expression
33
in developing hippocampal neurons (Kim et al., 2011). DHA and synaptamide appear to target
the same transcriptional activity, since both promote the expression of similar specific synaptic
proteins raising the possibility that their effect on neuritogenesis and synaptogenesis is
interdependent.
It is still early to claim that the neuritogenic and synaptogenic effects of DHA are mediated by
synaptamide. Although studies show the increase in synaptic puncta and neurite length with
synaptamide administration, the same are observed with DHA supplementation, albeit at a
higher concentration (Kim et al., 2011) (Rashid et al., 2013). The concentration difference
between the two compounds could be explained in two ways – one, to consider synaptamide
being synthesized from DHA; two, that synaptamide can more efficiently cross the membrane
than DHA hence requiring less concentration. Once in the cellular compartment, synaptamide
can liberate free DHA through hydrolysis which can be accounted for the functional effects
seen. Given the role of DHA in neurogenesis and less efficient transport of non-esterified DHA
across membranes, the notion of synaptamide delivering DHA which can be incorporated into
synaptic membranes appears to be a possibility. To begin to gather evidence for or against this
hypothesis, further studies of synaptamide are required. Some of the questions that need to be
answered in these studies are:
What is the biosynthetic pathway of synaptamide?
Is synaptamide synthesized in neurons or glia?
Can neurons take up synaptamide?
Is synaptamide synthesized ubiquitously or is it specific to specific brain regions?
Does any physiological response trigger the synthesis of synaptamide?
34
Is synaptamide synthesis observed only during developmental stages?
In the studies described in later sections of this dissertation, we attempted to understand what
happens to synaptamide in the brain; and if neuritogenesis and synaptogenesis seen after DHA
and synaptamide administration are due to synthesis of synaptamide from DHA or incorporation
of DHA by synaptamide.
The studies in the present Dissertation involved:
Administration of synaptamide labeled with carbon-14 in either the DHA or ethanolamine
moiety to mice, followed by evaluation of brain uptake of radiolabel and chromatographic
analysis of its chemical form
Radiochemical studies in N27 cells (rat fetal mesencephalic immortalized cells) to
complement the in vivo studies
Evaluation of the effects of unlabeled synaptamide (and of DHA) on neuritogenesis in
N27 cells.
With these studies we attempted to address our specific aims:
1. To detect the biosynthesis of synaptamide from exogenous DHA.
2. To examine in vivo and in vitro uptake of synaptamide.
3. To determine the role of FAAH in synaptamide metabolism and its partitioning into
phospholipids.
4. To analyze and compare a functional effect of synaptamide with that of DHA.
35
B. RATIONALE
DHA is known to enhance cognition and memory by increasing synaptogenesis and
neuritogenesis in the hippocampus. Kim et al carried out several studies in primary hippocampal
and cortical neurons and concluded that the functional effects of DHA in these cultures are
mediated by its conversion into synaptamide, a derivative of DHA (Kim et al., 2011). We
employed an immortalized fetal mesencephalic cell line constituting 95% TH+ immortalized
dopaminergic cells: 1RCN27 cells (N27 cells), which are glial cell free, in our studies for the
following reasons: (1) Previous studies on synaptamide were carried in primary cell cultures
which comprise a mixture of various cell types. By using N27 cells, we wanted to document if
the synthesis of synaptamide from DHA can occur in the neuronal compartment as opposed to
the glial compartment which is where DHA is known to be synthesized. (2) We aimed to
determine whether synaptamide synthesis from DHA is specific to hippocampal and cortical
glutamatergic neurons. As DHA is known to support neuronal survival and differentiation in
dopaminergic neurons through its action on RXR (de Urquiza et al., 2000; Perlmann et al.,
2004), we wanted to investigate the possibility of synaptamide being responsible for these
effects. (3) As N27 cells were derived from fetal cells, the undifferentiated cells represent the
fetal ―dividing‖ nerve cells. These cells take on the properties of an adult neural cell with the
onset of differentiation (Clarkson et al., 1999). Using these cells in our experiments may provide
us with a preliminary insight about the importance of synaptamide (and DHA) supplementation
in developing as well as mature, adult neural cells. Hence, we used both undifferentiated and
differentiating cells in our studies.
36
Arachidonic acid is an omega-6 fatty acid which is incorporated in its ethanolamide,
anandamide. Synaptamide incorporates DHA, an omega-3 polyunsaturated acid making it
structurally similar to anandamide. Based on this structural similarity, studies performed by Kim
et al largely assumed that synaptamide is synthesized from DHA endogenously via a similar
biosynthetic pathway as anandamide and that the functional effects of synaptamide are
terminated by its hydrolysis into DHA by FAAH. The rationale behind each of our specific aims
can thus be justified based on the following arguments:
1. The existence of synaptamide precursors (phospholipid precursors) can substantiate the
notion that synaptamide biosynthesis maybe similar to that of anandamide. One study
reported the presence of these precursors in bovine brain homogenates (Bisogno T,
1999) but no study addressed this issue in in vitro conditions. Thus, in our studies, we
attempt to investigate the presence of machinery responsible for synaptamide synthesis
in N27 cells.
2. Previous work carried out in our lab as well as by our collaborators demonstrated that
the N-acylethanolamines of fatty acids are better taken up in vivo than the free fatty
acids themselves. This was shown to be the case with the omega-6 fatty acid,
arachidonic acid and its N-acylethanolamine, anandamide (Glaser et al., 2006; Hu et al.,
In press), as well as with the shorter-chain, saturated fatty acid, myristic acid (C14:0)
and its N-acylethanolamine, myristoylethanolamide (Hu, 2016), which is an important
signaling molecule in plant species. The structural similarity between arachidonic acid
and DHA; and anandamide and synaptamide makes it reasonable to assume that in vivo
synaptamide uptake is higher than that of DHA. This may explain the potent activity of
37
synaptamide in the experiments of Kim and co-workers (Kim et al., 2011), but there is no
evidence for this phenomenon in vitro. Hence, we attempted to evaluate the differences
between the uptake of free fatty acid and its corresponding ethanolamide in in vitro
conditions and compare it with in vivo uptake.
3. The second major assumption by Kim and co-workers (Kim et al., 2011; Rashid et al.,
2013) is that FAAH metabolizes synaptamide to DHA and ethanolamine, terminating its
functional effects. However, the substrate preference of FAAH to N-acylethanolamines
with very long acyl chains (>C20) has not yet been demonstrated. One study indirectly
looked at FAAH‘s substrate preference towards synaptamide (Bisogno T, 1999);
nevertheless no definitive studies appear to have been conducted. Since evidence in
support of this assumption was lacking we investigated the role of FAAH on
synaptamide metabolism.
4. The observation that synaptamide brings about neuritogenesis and synaptogenesis in
primary hippocampal and cortical cells at a lower concentration than DHA prompted Kim
and co-workers to hypothesize that synaptamide mediates the functional effect of DHA
(Kim et al., 2011). We attempted to validate and explain this functional effect of DHA and
exogenous synaptamide using a secondary neuronal cell line consisting of immortalized
fetal mesencephalic N27 cells.
38
C. MATERIALS AND GENERAL METHODS
C.1. ANIMALS
Male Swiss Webster mice (Charles River Laboratories, Cambridge, MA) weighing 25 ~30 g
were used for all in vivo studies. Mice were maintained at the animal facility of Division of
Laboratory Animal Medicine (DLAM) on 12 hour alternating light and dark period, with access to
food and water ad libitum. Mice were treated in compliance with NIH guidelines for the use of
laboratory animals and according to a protocol approved by the Institutional Animal Care and
Use Committee (IACUC).
C.2. RADIOACTIVE COMPOUNDS
[14C]Arachidonic acid, [14C]ethanolamine and [14C-ethanolamine]anandamide were purchased
from American Radiolabeled Chemicals. [3H-ethanolamine]anandamide was obtained from
Moravek Pharmaceuticals. [14C]Docosahexaenoic acid was obtained from both American
Radiolabeled Chemicals and Moravek Pharmaceuticals. [14C-ethanolamine] synaptamide and
[14C-docosahexaenoyl]synaptamide were synthesized by Dr. Richard Duclos, Jr. in our lab.
C.3. CHEMICALS
RPMI 1640 with L-Glutamine (InVitrogen), Trypsin, Fetal bovine Serum, penicillin-streptomycin,
N1-supplement (Sigma-Aldrich), Dibutyryladenosine 3′,5′-byclic monophosphate sodium salt
(Sigma-Aldrich), dehydroepiandosterone, α-tocopherol (Sigma-Aldrich), non-radioactive
39
docosahexaenoic acid (Nu-Chek Prep), non-radioactive synaptamide (synthesized by Dr.
Duclos in our lab), paraformaldehyde, 10% gelatin solution, DAPI Fluoromount-G
(southernbiotech; 0100-20). FAAH inhibitor PF3845 was provided by Dr. Duclos. TRIS, bovine
serum albumin were obtained from Sigma-Aldrich. Chloroform, acetone and methanol were
obtained from (Fisher scientific). Authentic phospholipid standards: 1-palmitoyl-2-hydroxy-sn-
glycero-3-phosphoethanolamine (856705P), L-α-phosphatidylinositol (840044P), 1-palmitoyl-2-
hydroxy-sn-glycero-3-phosphocholine (855675P), 1,2-dipalmitoyl-sn-glycero-3-phospho-L-
serine (840037P), 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (850705P) and 1,2-
dipalmitoyl-sn-glycero-3-phosphocholine (850355P) were procured from Avanti Polar Lipids.
Solvable and Ultima Gold™ XR, high flash-point liquid scintillation counter cocktail (Perkin
Elmer Las Inc) are used for dissolving the isolated tissue and quantification of counts
respectively.
C.4. EQUIPMENT
LS6500 Multi-Purpose Scintillation Counter (by beckman Coulter), cyclone® Plus Storage
Phosphor Imager, Resolution Storage Phosphor Screen and the OptiQuant software (by Perkin
Elmer Las Inc).TLCs were performed using preabsorbent silica gel G plates (Analtech, Newark,
DE, USA) and silica gel 60 F254 plates (EMD Millipore). Olympus CX-51 Fluorescent microscope
was used to get images for neurite analysis and the software BIOQUANT was used to acquire
images.
C.5. N27 CELL CULTURE
40
The N27 cell line is an immortalized fetal rat mesencephalic cell line that produces dopamine
and expresses tyrosine hydroxylase and dopamine transporter. It was created by transfecting
fetal rat mesencephalon with a plasmid vector carrying LTa gene from SV40 virus (Adams et al.,
1996).
N27 cells (obtained as a gift from Dr. Freed‘s lab, University of Colorado; kindly donated by Dr.
Loring) were grown at 37°C in 5% CO2 in RPMI 1640 with L-glutamine supplemented with 10%
Fetal bovine Serum (FCS) and 1% penicillin-streptomycin. bells (between 8-20 passages) were
grown in 75 cm2 (T-75) cell culture flasks and for passaging, the confluent cells are separated
using 1.4 ul of trypsin. The cells were re-suspended in 10 ml fresh medium and seeded at a
density of 200,000 - 300,000 cells/well in 6 well plates at least 24 hours before experimental
treatment. For morphological studies cells are seeded on cover slips coated with 1 mg/ml
gelatin solution overnight inside 6-well plates (cell Treat). 24 hours following plating, the medium
was replaced with fresh medium supplemented with differentiating agents, Dibutyryladenosine
3′,5′-byclic monophosphate (dibutyryl cyclic AMP; 2mM) and dehydroepiandosterone (60 µg/ml)
and either DHA or synaptamide and the cells were allowed to differentiate for 72 hours. DHA
and synaptamide are complexed with fatty acid free BSA in the presence of α-tocopherol before
supplementation. 10 mM Stock solutions of DHA and synaptamide with α-tocopherol were made
in ethanol and aliquots for further dilution were stored at -80°C. Subsequent dilutions are made
at room temperature in 1% BSA; 20 µl of this diluted mixture is added to the culture medium
along with differentiating agent. The final concentrations of BSA and α-tocopherol in the culture
medium are 0.01% and 40 µM. The final concentrations of DHA and synaptamide in the culture
medium are 1 nM, 10 nM, 100 nM, 1 µM and 10 µM. After 72 hours, differentiation is stopped by
removing the medium and washing the cells with thrice with 1X PBS. cells grown on cover slips
41
are then fixed with ice cold 4% paraformaldehyde for 20 minutes in the fume hood followed by
three washed with 1X PBS to wash out most paraformaldehyde. The cover slips are then
mounted immediately for neurite analysis. Neurite analysis was performed on 70 randomly
selected cells and 80 randomly selected neurites from them.
C.6. STATISTICAL ANALYSIS
Statistical analysis in most experiments is performed using student‘s T test. For neurite analysis,
one-way ANOVA was used to compare the differences between the functional effects caused
various doses of either synaptamide or DHA. Post hoc tests were performed using post hoc T
test with Bonferroni‘s correction
42
D. SYNTHESIS OF SYNAPTAMIDE
D.1. INTRODUCTION
Endogenous or exogenous free fatty acids are able to undergo incorporation into complex lipids
only after they have been activated into their respective acyl-CoAs (Yan et al., 2015). Formation
of acyl-CoA utilizes ATP, coenzyme A and Mg2+ and ―fixes‖ the fatty acid inside the cell (Yan et
al., 2015). The conversion of fatty acid into fatty acid acyl-CoA is a two-step process: the fatty
acid first forms an intermediate with ATP (fatty acyl-AMP) which reacts with coenzyme A to form
fatty acyl-CoA (Mashek et al., 2007). The enzymes responsible for this conversion are acyl-CoA
synthases. Formation of arachidonyl-CoA and docosahexaenoyl-CoA require ASCL4 and
ASCL6 respectively (Kang et al., 1997; Marszalek et al., 2005a). Formation of an acyl-CoA is
not only an important step in its incorporation into phospholipids; it is also necessary for the
biosynthesis of the phospholipid precursors of the fatty acid‘s corresponding N-
acylethanolamine.
N-acylethanolamines (NAEs) are bioactive derivatives of fatty acids of various chain lengths that
have been identified in the mammalian brain. N-palmitoyl-, N-stearoyl- and N-oleoyl-, N-
linoleoyl-, N-linolenoyl-, N-dihomo-γ-linolenoyl- and N-arachidonoyl ethanolamine are some of
the most prominent N-acylethanolamines in the brain (Mechoulam et al., 1998). Many of these
N-acylethanolamines have specific functional roles in brain as well as other peripheral tissues –
N-palmitoylethanolamine has analgesic and anti-inflammatory functions whereas N-
oleoylethanolamine has a potent role in regulation of feeding (Freund T F, 2003). Anandamide
has a well-known role in the regulation of Cody temperature, locomotion, feeding and the
43
perception of pain, anxiety and fear (Walker et al., 1999),(Williams et al., 1999). The existence
of N-docosahexaenoylethanolamine (synaptamide) has been known for a while but its
―bioactive‖ role has only been recently postulated and in the last decade there has been some
focus on elucidating the possible functions of synaptamide.
Several biosynthetic pathways for the synthesis of N-acylethanolamines have been proposed.
While the most widely known and accepted pathway involves the production of N-
acylethanolamines from their phospholipid precursors: N-acylphosphatidylethanolamines
(NaPEs) by the action of the enzyme phospholipase D (Schmid H, 1996), there are several
alternate pathways that contribute to the N-acylethanolamine synthesis. These are
phospholipase D independent synthetic pathways – N-acylphosphatidylethanolamines can
either be cleaved by phospholipase C (PLC) or α/β-hydrolase 4 (ACh4; a serine hydrolase) into
phospho-N-acylethanolamines (pNAEs) or glycerophospho-N-acylethanolamine (GP-NAE)
respectively. The pNAEs are dephosphorylated by phosphatases and GP-NAEs are cleaved by
phosphodiesterases into the corresponding N-acylethanolamines. These phospholipase D
independent pathways serve as ―back up‖ methods for the synthesis of these biomolecules (Liu
et al., 2008).
Anandamide is the most studied N-acylethanolamine. Its implication in many physiological
effects makes it an attractive subject. Although synaptamide is a structural analog of
anandamide, it is not an endocannabinoid as it has very poor affinity for cannabinoid receptors –
Ki~400 nM for synaptamide vs 38 nM for anandamide at porcine CB1 receptors (Sheskin et al.,
1997) and Ki~12.2 μM for synaptamide vs 540 nM for anandamide at human CB1 receptors
(Felder et al., 1993) while at the CB2 receptors, the EC50 for synaptamide~9.8 nM vs 0.3 nM
44
for anandamide (Yang et al., 2011). As the two N-acylethanolamines are structural similar, the
information available about anandamide provides a resource to explore the fate of synaptamide.
One of the first proposed pathways for anandamide synthesis involves a simple, energy
independent condensation of free arachidonic acid and ethanolamine utilizing the enzyme
FAAH; in vitro evidence in support of this has been reported by several investigators (Devane W
A, 1994),(Kruszka et al., 1994). The apparent Km values of arachidonic acid and ethanolamine
for this reaction are ~100 μM and ~50 mM respectively (Ueda N, 1995). These values are much
higher than their physiological levels, so that formation of anandamide by reversal of the FAAH
reactions seems improbably (Siguira T, 1996). In fact, although this synthesis can be
demonstrated in vitro, evidence of this process in vivo is lacking. The most widely accepted
pathway involves the synthesis of anandamide from its phospholipid precursor, N-
arachidonoylphosphatidylethanolamine (NAPE; formed by the transacylase mediated N-
acylation of phosphatidylethanolamine). NAPE is then cleaved by an N-
arachidonoylphosphatidylethanolamine specific phospholipase D enzyme to release
anandamide and phosphatidic acid (Schmid H, 1996).
The first evidence of synaptamide synthesis was observed while investigating the synthesis of
fatty acyl ethanolamides from various long chain fatty acids (Devane W A, 1994). Synaptamide
synthesis was only mentioned, not stressed in this paper, as its levels were far less than those
of anandamide in bovine brain homogenates (Devane W A, 1994). The synthesis of
synaptamide from DHA was reported in hippocampal neurons (Cao et al., 2009), cortical
neurons and neural stem cells (Rashid et al., 2013); however, the mechanism of its synthesis
and the presence of its precursors in neural tissue was not characterized. There is no
45
information yet regarding the site of synaptamide synthesis – whether it takes place in neurons
or glia -- or on the conditions required for its synthesis. This chapter attempts to address some
of the gaps in knowledge of synaptamide biosynthesis.
D.2. METHOD
D.2.1. Synthesis of N-acylethanolamines in mouse brain homogenates
We used a protocol developed by Ueda et. al (Ueda N, 1995), and modified it by involving the
addition of other co-factors to study the synthesis of N-acylethanolamines (anandamide and
synaptamide) from their corresponding fatty acids. Mouse brains from male SW mice were
obtained after euthanasia by cervical dislocation and 20 mg/ml homogenates were prepared
fresh with tris-magnesium-EDTA containing 3% bovine serum albumin. Protease and
phosphatase inhibitors were added to minimize enzymatic activity and homogenates were kept
on ice until radiolabelled substrate was added. The various reagents added were: ATP (1 mM);
PE (0.1 mM); ethanolamine (10 mM) and coenzyme A (1 mM). Incubations were started by
addition of 20 µM [14C]arachidonic acid or [14C]DHA to the homogenate and the mixture was
incubated at 37ºC for 60 minutes. The incubation mixture was vortexed every 5 minutes to
ensure uniform tissue contact with the added reagents. After termination of incubations, the
tissue lipids were extracted using the ―Folch‖ method. An aliquot of the tissue chloroform extract
collected was used to count total radioactivity using the liquid scintillation counter. Another
aliquot was spotted on a TLC plate along with standards and run in the chloroform/methanol/
ammonia (60:30:1) solvent system. After drying, the plates were apposed to phosphor screens
for 24 hours which were then scanned with the cyclone Plus Imager to obtain autoradiographs.
46
Autoradiograph intensity was analyzed using the software OptiQuant. An average of four
experiments was considered for analysis.
D.2.2. Synthesis of N-acylethanolamines in N27 cells
N27 cells (both undifferentiated and differentiating cells) were incubated with 200 nM
[14C]arachidonic acid or [14C]DHA. After a set time, the medium was removed, and cell pellets
were harvested for analysis of radioactive lipids. An average of three experiments was
considered for analysis.
D.2.3. Lipid extraction
We followed the procedure of Folch et al. (Folch, Lees et al. 1957). 200 μl of the extraction
mixture (chloroform/methanol, 2:1) was added directly to the cell pellet or to the brain
homogenate and incubated at room temperature for 20 minutes. The suspension was sonicated
on ice twice (30 seconds each time) and was centrifuged at 14000 rpm for 15 minutes. The
supernatant was transferred to a tube with 0.9% NaCl (40 μl). 100 μl of chloroform was added to
cell debris for sonication and the suspension was centrifuged again for 15 minutes at 14000
rpm. The supernatants were mixed, vortexed and centrifuged again to separate the organic and
aqueous layers. Organic and aqueous phases were collected into separate tubes. Aliquots of
the aqueous and organic (chloroform) layers were assayed for total radioactivity by scintillation
counting and the remainder of the chloroform layer was stored at -80ºC until TLC analysis.
D.2.4. Radio-TLC analysis
47
Chloroform extracts were subjected to one-dimensional thin-layer chromatography (TLC) on a
preabsorbent 20 X 10 cm silica gel G plate for about 150 min, using a mobile phase containing
chloroform/methanol/ammonia (60:30:1 v/v). [14C]arachidonic acid and [14C]anandamide; and
[14C]DHA and [14C]synaptamide standards were used to identify corresponding spots for
anandamide and synaptamide. Quantification of radioactive hot spots was performed using
Optiquant software (Version 5). The accumulation of radioactivity on a radio-TLC is shown as
intensity (in DLU/mm2) with the different levels of signal intensity reflecting the different amounts
of radioactivity. The autoradiograph was divided into lanes and the intensity of spots (in
DLU/mm2) in each lane was used for analysis. Image analysis was performed after subtracting
the background from the entire lane.
D.3. RESULTS
D.3.1. Synaptamide is synthesized in mouse brain homogenates
TLC analysis of brain homogenates demonstrated the synthesis of N-acylethanolamine from the
radiolabeled fatty acid (fig D.1).
48
Fig D.1: Representative TLC used for the quantification of NAE synthesis. NAE synthesis ex-vivo
from 20 µM [14
C]arachidonic acid (A) and [14
C]docosahexaenoic acid (B). Legend (from left to
right): no added cofactors; added ATP (1 mM), PE (0.1 mM), EA (10 mM) and CoA (1 mM); no
added PE; no added EA; no added CoA; and no added ATP. (PE: phosphatidylethanolamine; EA:
ethanolamine; CoA: Co-enzyme A; ATP: Adenosine triphosphate)
We observed that even when no exogenous co-factors were added, there was some conversion
of the added free fatty acid into the corresponding ethanolamide. This can probably be
attributed to the presence of endogenous co-factors. As documented in fig D.1B, incubation of
brain homogenates with DHA resulted in the synthesis of synaptamide, possibly from a
corresponding N-docosahexaenoylphosphatidylethanolamine (NDPE) – PLD system. Addition
of all co-factors – phosphatidylethanolamine, ethanolamine, coenzyme A and ATP increased
anandamide and synaptamide synthesis (fig D.2). The absence of any one factor decreased
synaptamide synthesis but not to be low the endogenous baseline levels (fig D.2). This pattern
was consistent and reproducible thus confirming that all precursors play an important role in
synaptamide synthesis.
FIG D.1A FIG D.1B
49
Fig D.2: Quantification of TLCs of brain lipid extracts to determine NAE synthesis – Anandamide
from [14
C]arachidonic acid (A), and synaptamide from [14
C]DHA. (B). Data is expressed as an
average of four experiments in terms of DLU/mm2. Error bars represent SEM. Legend (from left to
right): no added cofactors; added ATP (1 mM), PE (0.1 mM), EA (10 mM) and CoA (1 mM); no
added PE; no added EA; no added CoA; and no added ATP. (PE: phosphatidylethanolamine; EA:
ethanolamine; CoA: coenzyme A; ATP: Adenosine triphosphate)
The synthesis of synaptamide was least in homogenate incubations without coenzyme A and
ATP (fig D.2D) indicating that the presence of adequate concentrations of these two compounds
is the most important requirement for synaptamide synthesis; they are needed for production of
the coenzyme A thioester of DHA.
D.3.2. In vitro synaptamide synthesis
FIG D.2A FIG D.2B
50
Lipid extracts of N27 (undifferentiated and differentiating) cells incubated with [14C]arachidonic
acid or [14C]DHA displayed a time dependent increase in radioactive counts (fig D.3).
Fig D.3: % Radioactivity in chloroform extracts of N27 cells as determined from scintillation
counts. N27 cells incubated with 200 nM [14
C]arachidonic acid (A) and [14
C]DHA (B). Data is
expressed in terms of % radioactivity in cell extracts. N=3; error bars represent SEM.
The time dependent increase in radioactivity of cell extracts could represent increased uptake of
[14C]arachidonic acid which in turn results in increased synthesis of its metabolites. To confirm
this, the chloroform extracts were subjected to TLC analysis (fig D.4).
FIG D.3A FIG D.3B
51
Fig D.4: Representative TLC images NAE synthesis from radiolabeled free fatty acid. Anandamide
synthesis in undifferentiated (A) and differentiating (B) N27 cells incubated with 200 nM
[14
C]arachidonic acid. Synaptamide synthesis in undifferentiated (C) and differentiating (D) N27
cells incubated with 200 nM [14
C]DHA.
TLC analysis revealed no discernible free [14C]arachidonic acid or [14C]DHA in cell extracts and
that these exogenous fatty acids were converted into metabolites by the cells. To confirm the
observation from scintillation counting of chloroform extracts, we quantified the hot spots on
TLC image performing lane analysis. As evident from the TLC, there was a clear quantifiable
time dependent increase in total radioactivity in each lane (fig D.5A and B) which correlates well
with the scintillation counts (fig D.3A and B). Autoradiographs of the TLC images have a
FIG D.4A FIG D.4B FIG D.4C FIG D.4D
52
―smudgy‖ appearance just below the actual hot spot, and this can be accounted to air-oxidation
of the standard as it runs along the silica plate in the mobile phase or as it dries out.
Fig D.5: TLC Quantification of chloroform extracts of cells incubated with radiolabelled fatty acids.
(A) [14
C]arachidonic acid and (B) [14
C]DHA. (C) Anandamide synthesis in N27 cells from 200 nM
FIG D.5A FIG D.5B
FIG D.5C
53
[14
C]arachidonic acid. Data is expressed in terms of % DLU/mm2 normalized to the background.
N=3; error bars represent SD.
Anandamide synthesis was evident from the TLC (fig D.4) as its RF corresponded to that of the
standard. The extent of synthesis suggests time dependence, possibly contributing to the time
dependent radiolabel accumulation in cell extracts. Quantification of hot spots shows that the
synthesis of anandamide occurs to a similar extent in both undifferentiated and differentiating
cells (fig D.5C). This suggests that the decrease in total radiolabel accumulation in
differentiating cells does not reflect decreased anandamide synthesis but is caused by
decreased incorporation of the label into phospholipids.
In contrast to the situation with incubations containing [14C]arachidonic acid, synaptamide
synthesis from [14C]DHA was not observed in N27 cells. TLC analysis of cells treated with
[14C]DHA did not show a corresponding radioactive spot with the synaptamide standard (fig
D.4C and D). In order to confirm this, we co-spotted radioactive cell extracts (incubated with
[14C]DHA) and non-radioactive synaptamide and repeated the TLC analysis. Once the TLCs
were run and autoradiograph was recorded, we sprayed the TLC plates with copper
sulfate/phosphoric acid reagent, and the plates were charred in a hot oven. Charred spots
corresponding to the non-radioactive synaptamide standard were not observed, but several
other spots were revealed (fig D.6).
54
Fig D.6: Autoradiograph (right) and charred (left) TLC image representative of synaptamide
synthesis in N27 cells incubated with 200 nM [14
C]DHA. Charred plates displayed non-radioactive
synaptamide (1) and various lipid classes DHA incorporates into: phospholipids (2, 3, and 4) and
other triglycerides (top). The autoradiograph displays lipid classes that incorporated [14
C]DHA.
D.4. DISCUSSION
N-acylphosphatidylethanolamines (NAPE’s) are the main phospholipid precursors for the
synthesis of N-acylethanolamines (Schmid H, 1996). via phospholipase D (Ueda et al., 2010).
Synthesis of N-acylphosphatidylethanolamine is the rate limiting step in N-acylethanolamine
formation and is usually mediated by the membrane associated enzyme, N-acyltransferase
(NAT). N-acyltransferase catalyzes the trans-acylation reaction (involving the transfer of the acyl
group in the sn-1 position) between a donor phospholipid (phosphatidylethanolamine,
1
2
3
4
COLD SYN
[14C] DHA
COLD SYN + [14C] DHA
COLD SYN
[14C] DHA
COLD SYN + [14C] DHA
55
phosphatidylcholine, sn-1-acyl-sn-2-lysophosphatidylethanolamine or sn-1-acyl-sn-2-
lysophosphatidylcholine) and phosphatidylethanolamine. N-acyltransferase activation is
triggered in the presence of high levels of intracellular calcium (represents ―demand‖ for the
synthesis of N-acylethanolamine). (Wang et al., 2009). Members of the HRAS-like tumor
suppressor family (H-rev107 family) were reported to possess phospholipase activity as well as
O-acyltransferase activity (referred to as PLA/AT) and are also able to synthesize N-
acylethanolamine precursors (Uyama et al., 2012). One out of the five members of this family,
Hrasls5 (PLA/AT-5) has an N-acyltransferase-like activity which is independent of intracellular
calcium – hence the name iNAT (Jin et al., 2007). N-acylphosphatidylethanolamines synthesis
from free fatty acids or fatty acyl Co-A was documented only in plants (McAndrew et al., 1998).
Since the conditions for anandamide synthesis are well documented, the same were used to
determine the synthesis of synaptamide. The precursor of anandamide synthesis is N-
arachidonoylphosphatidylethanolamine, formed from phospholipids incorporating arachidonic
acid. The formation of arachidonoyl-phospholipids be gins with the synthesis of arachidonoyl-
CoA which is an energy dependent reaction utilizing ATP and coenzyme A (Wilson D B, 1982).
The concentrations of exogenous docosahexaenoic acid (20µM), coenzyme A (1 mM) and ATP
(10 mM) were determined based on the apparent Km values of each factor listed in literature for
anandamide synthesis. Arachidonic acid (Km: 100 µM) utilizes coenzyme A (Km: 180 µM) and
ATP (Km: 0.5 mM) to form arachidonoyl-CoA (Wilson D B, 1982). Phosphatidylethanolamine
and ethanolamine (50 mM) were also added to determine if they play a role in increasing the
synthesis of synaptamide. Our results clearly show that the involvement of ATP and Coenzyme-
A play an important role in synaptamide synthesis. This is in agreement with the fact that
formation of docosahexaenoyl-CoA is the most important step for the synthesis of the
56
phospholipid precursor (N-docosahexaenoylphosphatidylethanolamine; NDPE) for the synthesis
of synaptamide.
While anandamide was synthesized from [14C]arachidonic acid in N27 cells, we were not able to
document the synthesis of synaptamide in N27 cells under the same conditions. This suggests
that the conditions required for the formation of docosahexaenoyl-CoA in vitro is different from
those required for the synthesis of arachidonoyl-CoA. It is also possible that N27 cells do not
express the required enzyme (ASCL6) for the synthesis of docosahexaenoyl-CoA: there are no
studies that demonstrated either the presence or absence of this enzyme in these cells. The
cells were incubated with the radiotracer for a maximum of 20 minutes. Another possibility for
the lack of synaptamide synthesis is that the incubation time may not be sufficient for its
synthesis from [14C]DHA. This is however unlikely as when cells were incubated with exogenous
synaptamide, it was rapidly metabolized (explained in detail in sections D and E) into
phospholipids. Therefore, if the rate of synthesis of synaptamide is slower than the rate of its
metabolism, intact synaptamide cannot be isolated or visualized.
D.5. CONCLUSION
Long-chain N-acylethanolamines (NAEs) and their precursors, N-
acylphosphatidylethanolamines (NAPEs), are trace constituents present in all cells and tissues.
Their cellular levels are tightly regulated. While saturated and monounsaturated N-
acylethanolamines represent the vast majority of cellular N-acylethanolamines, polyunsaturated
NAEs (especially arachidonoylethanolamide: anandamide) are produced ―on demand‖ and elicit
important physiological effects (Schmid, 2000). Docosahexaenoylethanolamide (synaptamide)
57
is reported to have bioactive properties and its mechanism of synthesis has not yet been
explored. The process of endocannabinoid biosynthesis remains enigmatic despite the
abundance of studies performed to characterize it; especially anandamide – owing to its
implication in a number of important physiological functions. Under previously established
protocol, we reproduced the synthesis of anandamide from arachidonic acid as well as
demonstrated the synthesis of synaptamide from DHA in crude brain homogenates but failed to
do so in vitro in N27 cells.
58
E. UPTAKE OF SYNAPTAMIDE – IN VITRO AND IN VIVO STUDIES
E.1. INTRODUCTION
Anandamide, N-arachidonoylethanolamine, is an endogenous ligand of the cannabinoid
receptors (CB1 and CB2) and of vanilloid receptors that functions as a neuromodulator and
affects a variety of physiological processes. Anandamide and other N-acylethanolamines are
particularly interesting as they are not synthesized and stored in vesicles like conventional
neurotransmitters but are synthesized ―on demand‖. The trigger for synthesis of anandamide is
elevation of intracellular calcium levels through cellular depolarization or receptor stimulation
(Rodriguez de Fonseca et al., 2005). Acylethanolamine action is terminated by enzymatic
hydrolysis.
Anandamide is released from the postsynaptic membrane when triggered by the calcium influx
that occurs by the activation of presynaptic glutamate receptors (Galligan, 2009). The
anandamide released can either act on postsynaptic cannabinoid receptors and inhibit
activation of excitatory stimulus or act on presynaptic cannabinoid receptors and inhibiting the
presynaptic stimulus. Although evidence supports a postsynaptic action of anandamide
(Giuffrida et al., 1999), its action on presynaptic receptors is dominant mainly because of the
abundance of CB1 receptors on the presynaptic terminals (Schlicker et al., 2001). Anandamide
hence is involved in ―retrograde signaling‖ where it causes a depolarization induced suppression
of inhibition (DSI) (Pitler et al., 1994; Wilson et al., 2001a; Wilson et al., 2001b) or depolarization
induced suppression of excitation (DSE) (Diana et al., 2004) in presynaptic terminals.
59
Clearly, in order for anandamide to act on the presynaptic terminals, it has to cross membranes
to reach the receptor. In addition, the enzyme responsible for anandamide hydrolysis, FAAH
(Fatty Acid Amide Hydrolase) is localized to the Endoplasmic Reticulum (Gulyas et al., 2004) so
anandamide will have to reach the intracellular compartment to be inactivated. This makes the
study of uptake and transport of anandamide an important issue. Several attempts have been
made to investigate the mechanism behind anandamide uptake and intracellular transport. The
first report of anandamide uptake in cells was published in 1993 (Deutsch et al., 1993) and
since then, many studies have explored the mechanism(s) by which it enters cells. One of the
first mechanisms proposed to explain anandamide transport was energy independent,
saturable, time- and temperature- dependent facilitative diffusion which was reported to be
bidirectional (Hillard C J, 1997). Other studies reported an unsaturable anandamide uptake in
certain cell lines (Fasia et al., 2003). This controversial observation prompted the re-
examination of anandamide transport and results were interpreted by different workers both in
support of as well as against the involvement of a specific transporter protein. FABPs (Fatty
Acid binding Proteins), in particular FABP5 and FABP7 are shown to be involved in the uptake
of anandamide along with other lipophilic compounds such as fatty acids and fatty acid amides
(Sanson et al., 2014). A variant of FAAH, FLAT (FAAH-like-anandamide-transporter) was also
shown to specifically mediate anandamide transmembrane transport (Chicca et al., 2012; Leung
et al., 2013) suggesting that multiple mechanisms play a role in intracellular anandamide
trafficking. The details of the transmembrane transport are not clear yet. There has been a lot of
argument in support of, as well as against, the existence of a specific anandamide
transmembrane transporter. However, it is notable that no such specific transporter has been
cloned. A recent review article by Dale G. Deutsch on anandamide trafficking reflects that unlike
classical neurotransmitters, anandamide does not require a protein to enable its transmembrane
60
transport, and that its intracellular trafficking is coupled to its internalization by fatty acid binding
proteins and subsequent hydrolysis by the enzyme fatty acid amide hydrolase (Deutsch, 2016).
Although much progress has been made in understanding these events, investigation of
diffusion and transport of highly lipophilic signaling molecules (such as anandamide) is fraught
with difficulties, and in vitro assays can suffer from artifacts and false negatives. Because of its
lipophilicity, the solubility of anandamide in aqueous media is extremely low and high non-
specific binding occurs to glass and plastic surfaces (Oddi et al., 2010), as well as to lipid
components of cells, tissue homogenates or membrane preparations used in experiments. This
makes the determination of the free concentration of anandamide impossible. Incorporation of
Bovine Serum Albumin (BSA) into assay media may minimize sticking to surfaces, but does not
alter the fact that the concentration of ―free‖ anandamide available for transport or enzymatic
hydrolysis is unknown. Thus kinetic parameters describing transport or metabolism are highly
dependent on experimental conditions. The concentration of BSA is thus an important
experimental variable. Albumin is a known carrier for a variety of hydrophobic compounds
including anandamide (Giuffrida et al., 2000). The concentration of albumin in human plasma is
~650 μM and that of anandamide is about 50 -200 nM making it possible for albumin to bind all
of the anandamide. It has been estimated that the free anandamide-to-bound anandamide ratio
in plasma is approximately 0.01% (Bojesen et al., 2003). The choice of BSA is hence important
to make sure that the non-specific binding of anandamide decreases while its uptake remains
significantly un-inhibited. Another important factor in determining anandamide uptake is
temperature. Many authors have reported a temperature dependent uptake of anandamide in –
cultured brain neurons (Di Marzo V, 1994), mouse N2A neuroblastoma and RCL-2H3 basophilic
leukemia cell lines (Jacobsson et al., 2001) and human keratinocytes (Oddi et al., 2005); the
61
optimal uptake of anandamide in all cases being observed at 37ºC. One strategy to discern the
―specific uptake‖ of anandamide is to determine its uptake at 4ºC (where most cellular activity is
stopped) and subtracting it from the uptake observed at 37ºC (Thors et al., 2006).
Anandamide clearance from the extracellular domain and hence its uptake is determined as a
combination of its association with membrane lipids and its transportation into the cells. This
cellular uptake proceeds until equilibrium is established between intra- and extracellular
compartments. Intracellular metabolism of anandamide is a rate limiting factor that governs the
establishment of equilibrium and hence anandamide uptake into the cell (Fowler, 2012). FAAH
is associated with metabolizing anandamide in the cell and hence is postulated by many authors
to be involved in driving anandamide uptake (Glaser et al., 2005). None of the groups
steadfastly claimed that FAAH is the sole reason for anandamide uptake as some authors
reported evidence for a FAAH independent uptake of anandamide (Beltramo et al., 1997;
Kathuria et al., 2003).
The subject of anandamide uptake and metabolism, although well characterized, is still a
subject of great controversy. However, the models used for this study can also be extended to
explore the uptake of synaptamide. As synaptamide is structurally similar to anandamide, many
aspects involved in its uptake can be compared against those of anandamide, thus fostering
better understanding of synaptamide uptake.
E.2. METHODS
62
E.2.1. Determination of synaptamide partitioning between Blood and brain compartments over
time.
1 μCi of [14C]synaptamide in 200 µl emulphor/ethanol/saline (1:1:18) was administered to Male
SW mice weighing 25-30g via tail vein injection. The solution for i.v. injection was prepared just
before starting each experiment, to limit the rearrangement or degradation of radiotracer.
Ethanol (the solvent in which the radiotracer was dissolved) was evaporated under a flow of
argon to protect against oxygen, after which the tracer was re-dissolved in
emulphor/ethanol/saline (1:1:18) vehicle. Before injecting into mice, 20 µl of the injection mixture
was assayed with the scintillation counter for the amount of radioactivity for the determination of
injected activity and the calculation of IA%/g values. After 0, 5, 15, 30 or 60 minutes mice were
euthanized by cervical dislocation, the brains removed immediately and placed in ice cold saline
for a few minutes to suspend enzymatic activity. Half the brain was weighed in scintillation vials,
and dissolved in ―Solvable‖ (tissue solubilizer). Scintillation cocktail was added to this mixture
and the radioactivity was assayed. Blood and urine samples were collected with fine tip transfer
pipettes when mice were euthanized, and weighed in scintillation vials. Blood samples were
bleached with hydrogen peroxide after dissolution and before addition of cocktail. Urine samples
were directly mixed with scintillation fluid. The counts per minute (CPM) and H# obtained from
the liquid scintillation counter were used to calculate the percent of injected activity per gram
(%IA/g) for the samples. The Blood/brain radioactivity concentration ratios were then calculated.
Two-tailed unpaired student's t test was used to make comparisons between percent IA/g
values with [14C]synaptamide, [14C]DHA and [14C]ethanolamine were made. Remaining half
brain and Blood samples were stored at -80ºC until TLC analysis. Data from three mice was
used for analysis.
63
E.2.2. Analysis of radiotracer uptake in vivo: microdissection studies
Male SW mice (25-30g) were used for these experiments. Each mouse was administered 0.1
μCi of [14C]radiotracers in 200 µl emulphor/ethanol/saline (1:1:18) via tail vein injection. After 15
minutes, mice were euthanized by cervical dislocation, the brains removed immediately, and
microdissected on a filter paper wetted with 0.9% saline, using the forceps method.
Hypothalamus, olfactory tubercle, frontal cortex, hippocampus, striatum, cerebellum, brain stem,
mid brain, thalamus and rest of the brain were separated, weighed in scintillation vials, and
dissolved in tissue solubilizer. Scintillation cocktail (2-5 mL) was then added to each vial, and
the radioactivity quantified using liquid scintillation. Blood and urine samples were treated as
described above (E.2.1). In view of the low radioactivities injected, samples were counted for 30
minutes and carefully corrected for background. Data from eight mice was used for analysis.
E.2.3. Determination of radiotracer uptake in vitro
In vitro uptake studies were performed by incubating N27 cells with various radiotracers:
[14C]arachidonic acid, [14C]anandamide, [14C]DHA and [14C]synaptamide. The uptake assay
procedure was based on the method used by Fowler et al. (Fowler et al., 2004), later modified
by Oddi et al (Oddi et al., 2010). N27 cells were passaged in RPMI 1640 with 10% FCS and
plated in 7 cm2 tissue culture dishes at a density of 106 cells/ dish. 24 hours following seeding,
the medium was replaced by 1X HBSS with 0.1% BSA and the cells were used for uptake
experiments. For differentiating N27 cells, the cells are seeded at the same density in the tissue
culture dishes, and after 24 hours, differentiating agents – Dibutyryladenosine 3′,5′-byclic
64
monophosphate (dibutyryl cyclic AMP; 2mM) and dehydroepiandosterone (60 µg/ml), were
added to the cells with fresh medium and allowed to differentiate for 48 hours. After 48 hours,
the medium was replaced with 1X HBSS with 0.1% BSA and the cells were used for uptake
experiments. The undifferentiated or differentiating cells were placed in a water bath maintained
at 37ºC. After about 10 minutes for equilibration at that temperature, the cells were incubated
with 200 nM radiotracer for 1, 2.5, 5 or 10 minutes. After the designated time, the reaction was
stopped by aspirating the HBSS into a scintillation vial and adding ice cold PBS with 1% BSA to
the petri dish and transferring it to ice. The cells were scraped off of the petri dish (in PBS) and
the suspension was centrifuged to get a cell pellet. The supernatant was discarded and the petri
dish was washed again with PBS with 1% BSA to recover any let over cells and this was added
to the cell pellet. The suspension was centrifuged again to recover the final cell pellet which was
kept on ice for lipid extraction. Control incubation was performed on ice and the cell pellet was
extracted using the same procedure as above.
For uptake studies with FAAH inhibited, the cells (both undifferentiated and differentiating cells)
were pre-incubated with 2 µM PF3845 in DMSO for 30 minutes at 37ºC before incubation with
the respective radiotracers. Data from three experiments for each condition was used for
analysis.
E.2.4. Lipid extraction from N27 cells
For lipid extraction of cell pellets from undifferentiated and differentiating cells, we followed the
procedure of Folch et al. (Folch, Lees et al. 1957). 200 μl of the extraction mixture
(chloroform/methanol, 2:1) was added directly to the cell pellet. The suspension was sonicated
on ice twice (30 seconds each time) and was centrifuged at 14000 rpm for 15 minutes. The
65
supernatant was transferred to a tube with 0.9% NaCl (40 μl). 100 μl of chloroform was added to
cell debris for sonication and the suspension was centrifuged again for 15 minutes at 14000
rpm. The supernatants were mixed, vortexed and centrifuged again to separate the organic and
aqueous layers. Organic and aqueous phases were collected into separate tubes. The aqueous
layer and an aliquot of the organic (chloroform) layer was used for scintillation counting to
determine the radioactivity portioned into each phase and the rest of the chloroform layer was
stored at -80°C until TLC analysis.
E.3. RESULTS
E.3.1. In vivo synaptamide uptake
E.3.1.1. Exogenous synaptamide enters the brain.
Synaptamide administered to mice via tail vein injections rapidly gets into the brain – within a
minute. The ratio of synaptamide in blood to brain decreases in a time dependent manner with
almost negligible synaptamide levels in blood one hour after injection (fig E.1A).
66
Fig E.1: (A) Blood to brain ratio of [14
C]synaptamide. Data expressed as the ratio of average %IA/g
radioactivity of synaptamide in Blood and brain, (C) Average %IA/g radioactivity in Blood and
brain over time. %IA/g given as mean ± SD; error bars represent SD. n=3 mice per time point.
The decrease in blood: brain synaptamide ratio could indicate that the brain takes up more
synaptamide from Blood over the course of time. Individual analysis of %IA/g in blood, brain and
urine indicate that the brain uptake of synaptamide is relatively constant over time, while its
presence in the blood decreased with time (fig E.1B and table E.1). This observation is
consistent with previous studies suggesting that the uptake of radiolabeled lipid (arachidonic
acid) is independent of cerebral blood flow (Chang et al., 1997).
TIME (MIN) 0 5 15 30 60
% IA/g in BRAIN 1.66 ± 0.29 1.79 ± 0.11 1.68 ± 0.12 1.96 ± 0.22 1.66 ± 0.38
% IA/g in BLOOD 15.90 ± 2.59 2.00 ± 0.25 1.23 ± 0.37 0.86 ± 0.12 0.56 ±0.07
% IA/g in URINE 6.42 ± 7.61 23.48 ± 23.48 47.46 ± 39.9 36.03 ± 16.37 36.18 ± 0.38
Fig E.1A Fig E.1B
67
Table E.1: Time-course of 14
C concentration (% IA/g) in Blood, brain and urine of mice after
administration of [14
C-ethanolamine]synaptamide via a tail vein. Data are the mean SD; error
bars represent SD. n=3 mice per time point.
E.3.1.2. Exogenous synaptamide is taken up differentially into different brain regions
Regional brain uptake and distribution experiments were performed in mice using
microdissection experiments. [14C-ethanolamine]synaptamide was used to perform these
studies. To our knowledge, our laboratory is the first to synthesize 14C labeled synaptamide and
to perform uptake and distribution studies.
Fig E.2: brain regional distribution of radiolabel 15 min after animals were injected intravenously
with 0.1μCi [14
C]synaptamide, 0.1μCi [14
C]DHA or 1μCi [14
C]ethanolamine. Values of %IA/g are the
68
mean s.d. with n = 3 for ethanolamine, and n = 8 for DHA and synaptamide. *p<0.05 when
compared to DHA.
The percent injected activity per gram for [14C]synaptamide after 15 minutes is shown in Fig E.2.
In order to ensure that the pattern seen is due to the uptake of intact synaptamide and rule out
the possibility of synaptamide breakdown, we administered another set of animals with
[14C]ethanolamine and [14C]DHA. The uptake pattern observed with these tracers is quite
different from that of synaptamide; thus confirming that synaptamide enters the brain as an
intact molecule.
E.3.1.3. [14C]Synaptamide distribution pattern in mouse brain is different from that of
[14C]anandamide.
The uptake of anandamide and arachidonic acid has been evaluated using autoradiography
technique (Hu et al., In press). The uptake pattern from autoradiography images is quantified in
terms of radioactive intensity (DLU/mm2). Although anandamide is a structural analog of
synaptamide; due to the difference in technique used and output unit, we cannot compare the
uptake of anandamide and synaptamide. The pattern of their distribution can however, be
compared. The distribution pattern of anandamide and synaptamide are different indicating a
different uptake profile and the possible implication that its functions are quite different than
those of anandamide (Fig E.3).
69
Fig E.3: The pattern of distribution of ethanolamine as a control as evaluated by autoradiography.
The uptake pattern is uniform throughout except in ventricles (A and B) (Hu et al., In press) reused
with permission. The pattern of distribution of Anandamide as observed with autoradiography(C
and D) (Hu et al., In press). The pattern of synaptamide is quite different to the pattern of
anandamide or ethanolamine (E and F).
0
0.5
1
1.5
2
2.5
3
3.5
4
Cortex hippocampus thalamus striatum
%IA
/g
[14C] Synaptamide Absolute Values
0
0.5
1
1.5
2
2.5
3
3.5
4
Cortex hippocampus thalamus striatum
[%IA
/g (
regi
on
) /
%IA
/g (
wh
ole
bra
in)]
[14C] Synaptamide Relative Values
Fig E.3A and B
Fig E.3C and D
Fig E.3E and F
70
E.3.1.4. [14C]Synaptamide uptake in vivo is higher than that of [14C]DHA.
Another notable observation made from autoradiography experiments was that the extent of
anandamide uptake is consistently higher than that of its corresponding acid, arachidonic acid
(table E.2). 36 days of exposure for various coronal brain slices revealed that the accumulation
of radioactivity produced with [14C]anandamide is 3 to 5 fold higher than that produced with
[14C]arachidonic acid (table E.2) (Hu et al., In press).
Absolute signal intensity (DLU/mm2)
(36 days exposure)
Signal intensity
ratios
brain regions
Wait time
after
injection
[14C]arachidonic
acid [14C]anandamide
[14C]anandamide /
[14C]arachidonic acid
Cortex
10min 1.97E+06 9.45E+06 4.79
100min 2.26E+06 1.10E+07 4.87
Hippocampus
10min 1.33E+06 5.84E+06 4.38
100min 1.52E+06 7.06E+06 4.65
Ventricular
epithelium
10min 3.74E+06 1.41E+07 3.76
100min 5.84E+06 1.91E+07 3.27
Thalamus
10min 1.89E+06 8.25E+06 4.36
100min 2.11E+06 9.92E+06 4.69
Striatum
10min 1.47E+06 6.66E+06 4.53
100min 1.76E+06 8.26E+06 4.70
71
Table E.2: Absolute signal intensity ratios of radiotracer accumulation in some brain regions after
36 days of exposure for various coronal brain slices. Accumulation of [14
C]anandamide is higher
than that of [14
C]arachidonic acid. Reused with permission from (Hu et al., In press)
Based on their structural similarity, we expected synaptamide and DHA to follow a similar
pattern. Indeed, we did find that brain synaptamide uptake was higher than that of DHA (table
E.3). The % uptake was evaluated using microdissection experiments and comparing the %IA/g
values obtained by scintillation counting.
%IA/g [14
C]DHEA %IA/g [14
C]DHA
%IA/g
[14
C]DHEA/[14
C]DHA
Hypothalamus 2.34 0.95 2
Olfactory tubercle 1.65 0.90 2
Hippocampus 1.18 0.67 2
Striatum 1.25 0.75 2
cerebellum 1.91 0.85 2
Brain stem 2.29 0.88 3
Cortex 1.47 0.95 2
Thalamus 1.19 0.83 1
Rest of the brain 1.15 0.59 2
Midbrain 2.95 0.93 3
whole brain 1.53 0.71 2
Blood 2.10 1.25 2
72
Table E.3: Ratios of % 14
C accumulation in various brain regions following the intravenous
administration of [14
C]DHA or [14
C]synaptamide in mice. N=8.
E.3.2. In vitro uptake studies
Previous in vitro studies demonstrate that a linear uptake trend for anandamide is observed
within the first 10 minutes of substrate addition (Fowler et al., 2004). Since little is known about
synaptamide uptake, the same conditions used to determine anandamide uptake were used to
study synaptamide uptake. We thus performed our uptake studies for 10 minutes – N27 cells
were incubated with 200 nM [14C-ethanolamine]anandamide for 1, 2.5, 5 and 10 minutes or [14C-
ethanolamine] synaptamide for 1, 2.5, 5, 10 and 20 minutes. The 20 minute time point was
chosen because in our preliminary studies we saw a slow but persistent accumulation of
radioactivity in the chloroform extracts over 10 minutes, and we wanted to determine whether
allowing the cells to incubate with synaptamide for a longer time point would yield the same
results as anandamide. To account for non-specific binding to culture plates, we performed
control incubation with cells on ice. In all cases, >90% of the radioactivity added to the
incubations was recovered. bell pellets were harvested and lipids were extracted in chloroform.
Hydrolysis of anandamide and synaptamide by the membrane enzymes is expected to release
[14C]ethanolamine which is water soluble and thus partitions into the aqueous phase and can be
easily quantified. The percentage of lipid uptake by the cells was determined by subtracting the
cell radiolabel uptake in control incubation from their uptake at 37ºC.
E.3.2.1. Anandamide and synaptamide have similar uptake profiles in N27 cells.
73
The scintillation counts from the aqueous phase of cell lipid extracts revealed a time dependent
accumulation of radioactivity – which indicates that [14C-ethanolamine]anandamide as well as
[14C-ethanolamine]synaptamide are hydrolyzed to release water soluble [14C]ethanolamine in a
time dependent manner (fig E.4A and D) in undifferentiated cells. The scintillation counts from
the chloroform extract also revealed a time dependent increase in radiotracer accumulation in
undifferentiated cells, but not in differentiating cells (fig E.4A, B, D and E).
Fig E.4: Comparison of uptake versus hydrolysis with [14
C]anandamide and [14
C]synaptamide in
N27 cells: [14
C]anandamide uptake and hydrolysis in (A) undifferentiated and (B) differentiating
N27 cells; (C) [14
C]anandamide uptake in undifferentiated and differentiated cells;
[14
C]synaptamide uptake and hydrolysis in (D) undifferentiated and (E) differentiating N27 cells;
Fig E.4A Fig E.4B Fig E.4C
Fig E.4E Fig E.4D Fig E.4F
74
(F) [14
C]synaptamide uptake in undifferentiated and differentiated cells. Data is expressed in terms
of % anandamide or synaptamide in cell lipid extract. Error bars represent SD; N=3. Statistical
analysis is done using student’s T test. *p<0.05 (comparison between uptake and hydrolysis at
each time point: A,B,D,E; comparison of uptake in undifferentiated and differentiating cells: C,F)
With both NAEs, uptake was significantly higher in undifferentiated cells than in differentiating
cells (fig E.4C and F).
The synaptamide uptake pattern was similar to that of anandamide, but its uptake was lower
than that of anandamide in both undifferentiated and differentiating cells (fig E.5 A and B). On
comparing the hydrolysis of both anandamide and synaptamide, in undifferentiated and
differentiating cells, the rate of hydrolysis of anandamide is significantly higher than that of
synaptamide (fig E.5 C and D; p<0.05). This suggests that the enzyme that hydrolyses N-
acylethanolamines hydrolyzes anandamide preferentially.
Fig E.5A Fig E.5B
75
Fig E.5: Comparison of uptake and hydrolysis with [14
C]anandamide versus [14
C]synaptamide in
N27 cells: [14
C]anandamide and [14
C]synaptamide uptake in (A) undifferentiated and (B)
differentiating cells. [14
C]anandamide and [14
C]synaptamide hydrolysis in (C) undifferentiated and
(D) differentiating cells. Data is expressed in terms of % anandamide or synaptamide in cell lipid
extract. Error bars represent SD; N=3. Statistical analysis is done using student’s T test *p<0.05
(comparison between anandamide and synaptamide uptake: A, B and hydrolysis: C, D)
E.3.2.2. Anandamide and synaptamide uptake in undifferentiated cells is regulated by their
hydrolysis.
Anandamide uptake has been a subject of great controversy. Anandamide transport can be
considered as a three step process – adsorption, transmembrane transport and desorption
(Glaser et al., 2005). It is probably that anandamide diffuses passively across the cell
membrane, and that metabolism by FAAH into arachidonic acid and ethanolamine inside the
cell then drives the further uptake of anandamide into the cell, as equilibrium in concentration
gradient cannot be attained. Since we observed a similar pattern of uptake with both
anandamide and synaptamide, a similar effect of FAAH is expected to occur with the uptake of
Fig E.5C Fig E.5D
76
both tracers. To determine the role of FAAH in determining the uptake pattern of anandamide
and synaptamide in N27 cells, we pre-incubated the cells with a specific FAAH inhibitor, PF3845
and carried out lipid extraction and TLC analyses.
Fig E.6: Comparison of [14
C]anandamide uptake and hydrolysis in N27 cells with or without
PF3845: [14
C]anandamide uptake and hydrolysis in (A) undifferentiated and (D) differentiating N27
cells pre-incubated with PF3845. [14
C]anandamide uptake and hydrolysis with or without PF3845
in undifferentiated (C and C) and differentiating (E and F) N27 cells. Data is expressed in terms of
% anandamide in cell lipid or aqueous extract. Error bars represent SD; N=3. Statistical analysis is
Fig E.6A Fig E.6B Fig E.6C
Fig E.6D Fig E.6E Fig E.6F
77
done using student’s T test *p<0.05 (comparison between anandamide uptake: B and hydrolysis:
C in N27 cells incubated with or without PF3845)
When cells were pre-incubated with FAAH inhibitor, there was a significant decrease in
anandamide uptake and hydrolysis in undifferentiated cells suggesting that hydrolysis by FAAH
is the major driving force for anandamide uptake (fig E.6A, B and C). In contrast, in
differentiating cells FAAH inhibition did not seem to have an effect on either uptake or hydrolysis
(fig E.6D, E and F). This suggests that on differentiation, the N27 cells develop a FAAH
independent mechanism for the uptake of anandamide.
The results for synaptamide uptake studies after pre-incubation with PF3845 were similar to
those observed with anandamide. In undifferentiated cells synaptamide uptake and hydrolysis
rates were significantly lower when FAAH was inhibited (fig E.7A, B and C), while in
differentiating cells, on FAAH inhibition, there was no significant difference in the uptake of
either ethanolamide (fig E.7D, E and F).
Fig E.7A Fig E.7B Fig E.7C
78
Fig E.7: Comparison of [14
C]synaptamide uptake and hydrolysis in N27 cells with or without
PF3845: [14
C]anandamide uptake and hydrolysis in (A) undifferentiated and (D) differentiating N27
cells pre-incubated with PF3845. [14
C]synaptamide uptake and hydrolysis with or without PF3845
in undifferentiated (B and C) and differentiating (E and F) N27 cells. Data is expressed in terms of
% anandamide in cell lipid or aqueous extract. Error bars represent SD; N=3. Statistical analysis is
done using student’s T test *p<0.05 (comparison between synaptamide uptake: B and hydrolysis:
C, F in N27 cells incubated with or without PF3845)
These experimental results indicate that FAAH, which is known to hydrolyze various N-
acylethanolamines, may also be responsible for hydrolyzing synaptamide into DHA and
ethanolamine. However, the inhibition by FAAH is only observed at longer time points for
synaptamide versus earlier time points for anandamide (fig E.6C vs E.7C; fig E.6C vs E.7C).
This indicates that FAAH has a greater ability to hydrolyze anandamide than it does
synaptamide. It was also found that FAAH does not completely inhibit synaptamide hydrolysis in
undifferentiated cells (fig E.7C) suggesting that FAAH may not be the sole enzyme that can
hydrolyze it.
Fig E.7D Fig E.7E Fig E.7F
79
E.3.2.3. Anandamide and synaptamide uptake into undifferentiated N27 cells is similar to that of
AA and DHA but not in differentiating cells.
As we noticed different uptake rates for AA and anandamide as well as DHA and synaptamide
in vivo, we undertook to compare uptake of the two ethanolamides in vitro. The uptake studies
were repeated with 200 nM [14C]arachidonic acid and [14C]DHA and the uptake studies in
undifferentiated and differentiating cells were compared (fig E.8).
FIG E.8A
FIG E.8B
FIG E.8C FIG E.8D
80
Fig E.8: Comparison of [14
C]arachidonic acid and [14
C]DHA uptake in N27 cells: (A)
[14
C]arachidonic acid and (B) [14
C]DHA uptake in undifferentiated and differentiating N27 cells.
Comparison of [14
C]arachidonic acid versus [14
C]DHA uptake in undifferentiated (C) and
differentiating (D) N27 cells. Data is expressed in terms of % AA and DHA in cell lipid extract. Error
bars represent SD; N=3. Statistical analysis is done using student’s T test *p<0.05 (comparison
between uptake in undifferentiated and differentiating cells: A, B; and comparison between AA
and DHA uptake: C in undifferentiated cells)
Uptake of arachidonic acid and DHA, in both undifferentiated and differentiating cells, increased
with incubation time, although the time dependent increase was greater in undifferentiated cells
(fig E.8A and D). More arachidonic acid was taken up into undifferentiated and differentiating
cells than DHA (fig E.8C and D).
The uptake rates of arachidonic acid and DHA were similar to those of anandamide and
synaptamide in undifferentiated cells (fig E.9A and B). In differentiating cells, however, we
observed that the uptake rates of arachidonic acid and DHA were higher than those of
anandamide and synaptamide (fig E.9C and D) unlike our results in vivo.
FIG E.9A FIG E.9B
81
Fig E.9: Comparison of NAE uptake versus their corresponding fatty acid uptake in N27 cells: (A
and B) [14
C]arachidonic acid and [14
C]anandamide uptake and (C and D) [14
C]DHA and
[14
C]synaptamide in (A and C) undifferentiated and (B and D) differentiating N27 cells. Data is
expressed in terms of % radiotracer in cell lipid extract. Error bars represent SD; N=3. Statistical
analysis is done using student’s T test *p<0.05 (comparison between uptake of AA and
anandamide: C; and DHA and synaptamide uptake: D in N27 cells)
E.4. DISCUSSION
Previous in vivo work using [14C]arachidonic acid and [14C]anandamide from our lab
demonstrate that the two tracers have similar uptake profiles in the brain. However, the
accumulation of radioactivity after intravenous administration of [14C]anandamide is 3 to 5 fold
higher than that produced when [14C]arachidonic acid is injected (Hu et al., In press). This
phenomenon suggests that arachidonic acid (a fatty acid with negative charge and higher
polarity than anandamide) does not penetrate the Blood Brain Barrier as well as anandamide.
DHA (analogous to arachidonic acid) and synaptamide (analogous to anandamide) are both
expected to show a pattern similar to arachidonic acid and anandamide respectively, and the
FIG E.9C FIG E.9D
82
pattern indeed was similar with synaptamide uptake being at least 2 fold higher than that of
DHA.
We failed to replicate this observation in vitro. The uptake rates of arachidonic acid and DHA
were found to be higher than those of anandamide and synaptamide. This suggests that
specialized uptake mechanisms present in in vivo environments are absent in vitro. As several
studies established previously, in vitro fatty acid uptake is based on either the process of simple
diffusion or the presence of transporter proteins, or a combination of both(Qi et al., 2002). The in
vitro system is not a dynamic system – there is no Blood flow that provides the cells with a
concentration gradient or to remove the accumulated end points. This reinforces the idea that in
vitro uptake is a saturable process because without continuous clearance of end products, there
is a decrease in the corresponding uptake process.
Arachidonic acid and DHA exhibit higher uptakes in undifferentiated N27 cells than do their
ethanolamides; uptake of arachidonic acid being higher than that of DHA. Fatty acid uptake in
cells is driven by a combination of simple diffusion followed by their utilization (binding or
metabolism) (Kamp et al., 2006). Their utilization (and hence uptake) however depends on
membrane enzymes such as the phospholipases and acyl-CoAs. These enzymes partition the
available free fatty acids into different metabolic fates (Kalant D, 2004). Phospholipases,
specifically phospholipase A2s are responsible for releasing free arachidonic acid and DHA
from membrane phospholipids and Acyl Co-A synthase (ACS) enzymes are responsible for
―fixing‖ the fatty acid in the cell (Mashek et al., 2007). Consequently, the expression and activity
of the specific phospholipases and ACS enzymes determine the fatty acid uptake and
incorporation. The release of arachidonic acid and DHA from membrane phospholipids is
83
mediated by two separate phospholipases – calcium dependent cPLA2 is responsible for the
release of arachidonic acid and calcium independent (inducible) iPLA2 specifically releases
DHA (Strokin et al., 2003). The expression of these enzymes establishes the required
concentration gradient which drives the uptake of fatty acids. The released fatty acid is either
incorporated into other phospholipids or triglycerides, or is metabolized by a different set of
enzymes (LOX, COX, BYP450s, etc.) which generate second messengers that can start a
cascade of cellular functions (Rosa et al., 2009). This consumption of fatty acids maintains their
continuous influx simulating the ―dynamism‖ of in vivo environments.
According to the literature, 70% of PLA2 activity can be attributed to iPLA2 with its highest
activity recorded in striatum followed by hypothalamus and hippocampus while cPLA2 is
uniformly distributed all over the brain with a slightly higher activity in cerebellum than other
regions (Yang et al., 1999; Farooqui et al., 2004). Among various ACS enzymes, ACS6 is highly
expressed in brain tissues. This enzyme has a substrate specificity towards very long chain fatty
acids (>C20), specifically to DHA (Marszalek et al., 2005a). This indicates that DHA is
preferentially incorporated into the membrane and very little is available to be metabolized.
Arachidonic acid, on the other hand, is available for metabolism longer and hence its
incorporation into the membrane is slightly less than that of DHA. As a result, the turnover of
arachidonic acid is more rapid than that of DHA. We observed greater uptake of AA than DHA in
N27 cells. The relative expression of ACS6, PLA2 or COX and LOX in N27 cells is unknown and
hence information is lacking to explain the higher AA uptake.
Anandamide and synaptamide have similar uptake patterns in N27 cells as their corresponding
fatty acids. This is in contrast to what was observed in vivo where these ethanolamine
84
containing fatty acid acids are taken up to a higher extent. Metabolism of these compounds
inside the cells is expected to be the driving force of these compounds. However, compared to
anandamide, the cellular synaptamide uptake is less suggesting that its metabolism in N27 cells
is slower than that of anandamide. Anandamide is the preferred substrate of fatty acid amide
hydrolase (FAAH), a serine hydrolase which is usually associated with the inner cell membrane.
In some cell lines it has been shown that anandamide transport was indeed mediated by
modulating FAAH activity (Day et al., 2001; Fowler et al., 2004). We hypothesize that in N27
cells too, anandamide incorporates itself into the membrane by either a flip flop mechanism or
by coupling to a fatty acid binding protein making it accessible to the membrane associated
FAAH which then hydrolyses it into its corresponding fatty acid (AA) and ethanolamine, thus
creating a concentration gradient that drives the uptake of more anandamide into the cell. To
confirm that it is indeed FAAH that is hydrolyzing synaptamide influencing its uptake, the cells
were pre-incubated with a specific, irreversible FAAH inhibitor – PF3845. Clocking FAAH activity
markedly decreased synaptamide uptake as well as hydrolysis thus substantiating our
hypothesis. Combining the facts – synaptamide uptake being less than that of anandamide with
a further decrease in synaptamide uptake with PF3845 administration – it can be suggested that
synaptamide is a substrate for FAAH, albeit a poor substrate and thus explain its uptake pattern.
Differentiation of N27 cells was initiated by adding dibutrylcyclicAMP (dbcAMP) and
dehydroepiandosterone to the incubations. The uptake of fatty acids decreased considerably
with the onset of differentiation. DbcAMP is known to enhance the activity of iPLA2 and to thus
increase the release of DHA (Strokin et al., 2003). The decrease of DHA in the lipid extracts of
differentiating cells when compared to undifferentiated cells may be explained by the theory that
the amount of DHA released from the membrane is more rapid and consistent with addition of
85
dbcAMP than the amount of DHA being reincorporated. Increase in dbcAMP has no effect on
the activity of cPLA2 and the release of arachidonic acid (Strokin et al., 2003).
Before differentiation, N27 cells have a small nucleus and large cytoplasm – Undifferentiated
N27 cells are flat with a high cytoplasm-to-nucleus ratio. On differentiation, the cells become
more rounded, with decreased surface area and there is a large decrease in the cytoplasm -to-
nucleus ratio. As the rate of diffusion is expected to be proportional to the surface area of the
cell, it may be suggested that the amount of fatty acid diffusing into the differentiating cell is less
than that of the undifferentiated cell.
While the decreased uptake of DHA in differentiated cells may be explained based on above
observations, it is still unclear why arachidonic acid uptake is lower in differentiated cells. The
drop in surface area of the cells may contribute a little, but is not sufficient to explain the drastic
decrease.
The uptake of both arachidonic acid and of DHA in N27 cells decreased as the cells underwent
differentiation. Uncertain of whether this decrease is because of the changes in morphology of
the cells or because of the change in hydrolytic enzyme expression, experiments were repeated
after pre incubating differentiated cells with PF3845. Surprisingly there was no difference in the
uptake profiles when FAAH activity was blocked. It can thus be inferred that differentiation
modulates the activity of FAAH in a way that the uptake of the N-acylethanolamines becomes
FAAH independent. All the changes N27 cells undergo during differentiation have not yet been
documented, and further studies are required to explain the FAAH independent uptake process
in differentiating cells.
86
E.5. CONCLUSION
To our knowledge, ours is the first study that compares the uptake of free fatty acids and their
ethanolamine metabolites both in vitro and in vivo. Our previous observations that the brain
uptake of fatty acids is lower than that of their respective ethanolamides in vivo (Hu et al. in
press; Pandey et al. 2014) is shown in this dissertation to also be true for DHA and
synaptamide. However, this was not the case in N27 cells – where we found that both
compounds were taken up by the cells to the same extent until the initiation of differentiation
after which their uptake decreased considerably.
87
F. SYNAPTAMIDE METACOLISM: ROLE OF FAAH AND END FATE
F.1. INTRODUCTION
N-acylethanolamines that act on extracellular receptors are inactivated by uptake into the
postsynaptic cell followed by enzymatic hydrolysis (Gulyas et al., 2004). As anandamide is the
most extensively studied NAE, its metabolic pathway can be used to compare and understand
that of synaptamide. While the uptake of synaptamide is discussed in a previous chapter, this
chapter focuses on the second aspect of inactivation – enzymatic hydrolysis. It should be
noted, however, that in addition to hydrolysis to arachidonic acid and ethanolamine,
anandamide can undergo oxidation to bioactive metabolites by lipoxygenase (LOX),
cyclooxygenase (COX) and cytochrome P450‘s (Snider et al., 2009).
Fatty acid amide hydrolase (FAAH) is an intracellular serine hydrolase which hydrolyses N-
acylethanolamines (Ueda et al., 2000). FAAH is a membrane-bound protein, mainly found to be
associated with microsomal and mitochondrial subcellular fractions (Schmid et al., 1985). FAAH
can metabolize a number of other substrates in addition to anandamide such as – other fatty
acid amides (Cravatt et al., 1996) including oleamide (Maurelli s, 1995) and esters including 2-
arachidonoylglycerol (Goparaju et al., 1998). Structure analysis of FAAH revealed that it does
not have the classical Ser-His-Asp catalytic triad of serine hydrolase enzymes (Patricelli et al.,
1999). Instead, the catalytic site of FAAH contains two critical serine residues – ser217 and
ser241 – which when mutated, completely abolished its catalytic activity (Omeir et al., 1999).
Ser-241 of FAAH was identified as the catalytic nucleophile that can break the amide Cond of its
substrates (Patricelli et al., 1999).
88
DHA and AA compartmentalize differentially and independently (Rapoport, 2003) into
phospholipids. Published data indicates that DHA is predominantly incorporated into
phosphatidylethanolamine (PE) followed by phosphatidylserine (PS) and phosphatidylcholine
(PC) while AA is incorporated predominantly into phosphatidylinositol (PI) and
phosphatidylcholine (Martinez et al., 1998). Within these phospholipid classes the site of
acylation (the sn-1 or sn-2 position) depends on the particular phospholipases and
transacylases involved (Lamaziere et al., 2011). If synaptamide is hydrolyzed by FAAH into
DHA, it would incorporate into phospholipids in the order phosphatidylethanolamine >
phosphatidylserine > phosphatidylcholine.
As demonstrated by our in vivo studies, synaptamide is taken up into the brain. However, the
fate of synaptamide, once it crosses the BBB, is not known. Using in vitro and in vivo studies,
we aimed to investigate the chemical modifications and hence the intracellular end fate of
synaptamide.
F.2. METHODS
F.2.1. Microdissection studies with FAAH
Male SW mice (25-30g/mouse) were injected with 10 mg/kg PF3845 in DMSO (100 µl)
intraperitoneously 3 hours prior to radiotracer injection to inactivate the enzyme FAAH. The
mice were injected with [14C]synaptamide and were euthanized by cervical dislocation 15
89
minutes later. Brain dissection was then performed as described in section D.3.2. Data from
eight animals is used for analysis.
Another set of mice injected with [14C]synaptamide pretreated either with saline or PF3845 were
euthanized and the brain and blood samples collected were used for lipid extractions and TLC
analysis. Data from 5 animals is used for analysis
F.2.2. FAAH activity assay
FAAH activity was measured using a radiometric assay. We employed a modified version of the
protocol previously established by Deutsch and co-workers (Omeir et al., 1995). Mice were
euthanized by cervical dislocation and their brains were removed immediately. The whole brain
was homogenized using Tris Magnesium EDTA (TME) buffer with 2.5% Bovine Serum Albumin
(BSA) to make a final brain homogenate of various concentrations – 20 mg/ml, 10 mg/ml and
1.25 mg/ml. A portion of the brain homogenate prepared was pre-incubated with the selective
and irreversible FAAH inhibitor, PF3845 for 30 minutes at room temperature to validate that
these results are due to hydrolysis by FAAH. A solution of 0.2 mM PF3845 in DMSO was
prepared fresh before the start of each experiment. For incubations with inhibitor, 200 µl of the
tissue homogenates are taken into 1.5 ml microcentrifuge tube s and are supplemented with 2
µl of the prepared PF3845 solution. A stock solution of the substrate [14C-
ethanolamine]anandamide or [14C-ethanolamine]synaptamide (0.1μCi/100µl) was prepared and
kept on ice. 1µl of the prepared stock was added to the 200 µl of brain homogenates (of each
concentration) and incubated in the water bath at 37°C for 0, 15 or 30 minutes. This was
performed in triplicate. After the assigned time, the incubations were retrieved from the water
90
bath and the reaction stopped by the addition of a 1ml mixture of ice cold chloroform and
methanol (1:1) plus 250µl 2N Hydrochloric acid. Because the hydrolysis product,
[14C]ethanolamine is slightly basic, it partitions into the acid layer. The radioactivity of the acid
layer hence reflects the extent of hydrolysis. After 30 minutes, all samples were centrifuged for
12 minutes at 4⁰C and at 14,000 rpm. Following centrifugation, 200 μl of the upper, aqueous
layer was pipetted into liquid scintillation vials to which the Ultima Gold™ XR scintillation
cocktail was added and radioactivity measured using a liquid scintillation counter. The increase
in accumulation of radioactivity in the acid layer with time represents the rate of hydrolysis of
substrate by FAAH. Data from three experiments was used for analysis.
F.2.3. Competition binding assay
Reactions were carried out with mouse brain homogenates (1.25 mg/ml; 200µl) incubated with
different concentrations of non-radioactive anandamide or synaptamide or DHA and
ethanolamine over the range 1 μM – 1 mM. To these reactions, 2 µl radiolabelled anandamide
(0.5 µCi/200 µl stock) was added and the reactions were incubated at 37°C for 10 minutes. The
reactions were stopped by addition of 1ml mixture of ice cold chloroform and methanol (1:1)
plus 250 µl 2N Hydrochloric acid. Samples were then centrifuged at 4⁰C for 12 minutes at
14,000 rpm. 200 μl of the upper, aqueous layer was transferred to liquid scintillation vials to
which the Ultima Gold™ XR scintillation cocktail was added and radioactivity was measured
using a liquid scintillation counter. The radioactive counts thus obtained reflected the
percentage of total radioactivity released to the aqueous layer and were used to calculate the %
specific radioligand binding which was plotted against the log concentration of the inhibitor – in
91
this case, anandamide or synaptamide or DHA plus ethanolamine – using Graphpad Prism to
generate IC50 values in each case. Data from three experiments was used for analysis.
F.2.4. Lipid extraction
F.2.4.1. Lipid extraction from brain homogenates
Frozen brain samples were homogenized with chloroform and methanol (2:1 v/v) to make a
brain homogenate of 30 mg wet weight per ml. 250 µl of 40% urea and 250 µl of 5% sulfuric
acid were added to 500 µl of brain homogenate in a microcentrifuge tube and thoroughly
vortexed. When centrifuged, two layers were formed: a lower chloroform layer and an upper
aqueous layer with the protein deposit wedged in the middle. The aqueous and chloroform
layers were aspirated into separate tubes. To extract the lipids trapped under the protein
deposit, 200 µl of chloroform is added to it and was sonicated twice for 30 seconds each time.
Following vortexing and centrifugation the chloroform layer was added to that from the first
extraction. The aqueous layer was used for the quantification of aqueous metabolites and the
chloroform layer was stored at -80⁰C until used for TLC analysis.
Lipids from Blood samples were extracted similarly.
F.2.4.2. Lipid extraction from N27 cells
Lipid extraction of cell pellets from undifferentiated and differentiating cells was performed as
describe d in section C.3.4. Cell pellets were subjected to modified Folch extraction. Organic
and aqueous phases were collected. Radioactive counts in the aqueous layer and an aliquot of
92
the organic (chloroform) layer were determined by the liquid scintillation counter; and the rest of
the chloroform layer was stored at -80⁰C for TLC analysis.
F.2.5. Radio-TLC analysis
Chloroform extracts were dried under argon and then re-dissolved in 20 µl of chloroform. 1 µl of
the extract was used to determine the total amount of radioactivity. To be able to visualize the
metabolites efficiently with only a few days‘ exposure, a fraction of the extract containing at least
a 1000 CPM radioactivity had to be spotted on the silica gel plate. To avoid overloading the
plates, the minimum amount of extract containing at least 1000 cpm was spotted. Two types of
analyses were carried out – 1-dimensional TLC to separate the major lipid classes in the cell or
brain extracts, and 2-dimensional TLC to identify the different phospholipids into which the lipids
partitioned.
F.2.5.1. One-dimensional TLC
Chloroform extracts from brain, Blood or cells were spotted on a 20 X 10 cm silica gel 60G F254
plates and run for about 150 min, using a mobile phase containing chloroform–methanol–
ammonia (60:30:1 v/v). Fatty acid and fatty acid ethanolamide standards were used to identify
corresponding spots from reactions in which [14C]anandamide or [14C]synaptamide were used.
F.2.5.2. Two-dimensional TLC
93
Chloroform extracts from brain, Blood or cells were spotted on 10 X 10 cm silica gel 60G F254
plates and run in the first dimension for about 45 min in a mobile phase containing chloroform–
methanol–ammonia (65:35:5 v/v). The plates were allowed to dry thoroughly, turned at right
angles and then run in the second dimension for another 45 minutes in a mobile phase
containing chloroform-acetone-methanol-acetic acid-water (30:40:10:10:5). Authentic non-
radioactive phospholipid standards were visualized by charring.
Following development with the chosen solvent system in an air tight jar, the TLC plates were
air dried and then opposed to phosphor screens to produce autoradiograms. To confirm the
identities of radioactive spots, the TLC plates were either charred (to witness all standards and
metabolites – radioactive and non-radioactive) or sprayed with ninhydrin to visualize the
phospholipids with a free amino group. Iodine was also used to identify unsaturated
compounds.
F.3. RESULTS
F.3.1. Role of FAAH on synaptamide uptake in vivo
F.3.1.1. Effect of FAAH inhibitor on brain and Blood carbon-14 levels after [14C]synaptamide
Brain regional concentrations of 14C 15 min after intravenous injection of [14C]synaptamide to
control mice varied from 0.8 to 1.8 %IA/g in the order: brain stem>cerebellum>cortex,
hippocampus, striatum > rest of brain (Fig F.1). In mice pretreated with PF3845, the regional
differences were unchanged, but tissue radioactivity concentrations were 30-40% higher.
94
However, the increases did not achieve statistical significance (p<0.05) except for
hippocampus. Blood levels of radioactivity were not significantly different between control and
pretreated animals.
Fig F.1: brain regional concentrations of 14
C in animals euthanized 15 min after injection of 0.1μCi
[14
C] synaptamide (i.v.), with or without PF3458 pretreatment (10 mg/kg; i.p. 3h before radiotracer).
%IA/g values expressed as mean ± SD; error bars represent SD; n=8. Statistical analysis is
performed using Student’s T test. *p<0.05 when compared to synaptamide administration without
PF3845.
F.3.2. Role of FAAH on synaptamide uptake in vitro
F.3.2.1. Synaptamide undergoes hydrolysis by FAAH.
95
Figures F.2 and F.3 show the extent of hydrolysis of [14C-ethanolamine]anandamide and of [14C-
ethanolamine]synaptamide at 0, 15 and 30 min, using homogenate concentrations of 1.25, 10
and 20 mg/ml. Hydrolysis of both radiotracers increased with time and with homogenate
concentration, but the extent of hydrolysis was lower for synaptamide than for anandamide for
all conditions.
Fig F.2: Time dependent increase in radioactive counts in the aqueous layers of brain
homogenates treated with [14
C]anandamide and [14
C]synaptamide – (A) 1.25 mg/ml, (B) 10 mg/ml
and (C) 20 mg/ml. Data is expressed in terms of % radioactivity quantified in aqueous layers. Error
bars represent SD; N=3. Statistical analysis is performed using Student’s T test. *p<0.05
(comparison between anandamide and synaptamide hydrolysis)
Fig F .2A Fig F.2B Fig F.2C
96
Fig F.3: Time dependent increase in the hydrolysis of (A) [14
C]anandamide and (B)
[14
C]synaptamide in brain homogenates – 1.25 mg/ml, 10 mg/ml and 20 mg/ml. Data is expressed
in terms of % radioactivity quantified in aqueous layers. Error bars represent SD; N=3.
In order to confirm if FAAH hydrolyzed synaptamide, the assay was repeated after pre-
incubating the brain homogenates with 2 µM PF3845 (Ki for FAAH = 230 nM). With FAAH
inhibition anandamide hydrolysis decreased significantly at all conditions, while synaptamide
hydrolysis was inhibited only at longer time points and at the higher tissue concentration (fig
F.4).
Fig F.3A Fig F.3B
97
Fig F.4: Time dependent hydrolysis of [14
C]anandamide and [14
C]synaptamide in brain
homogenates treated with or without PF3845 – (A) 1.25 mg/ml, (B) 10 mg/ml and (C) 20 mg/ml.
Data is expressed in terms of % radiolabeled substrate hydrolyzed. Error bars represent SD; N=3.
Statistical analysis is performed using Student’s T test. *p<0.05 (comparison between anandamide
and synaptamide hydrolysis in the presence or absence of PF3845)
At time zero, there was no apparent effect of FAAH inhibition on anandamide or synaptamide
hydrolysis at any tissue concentrations.
F.3.2.2. The hydrolysis of synaptamide by FAAH is spontaneous as well as tissue mediated.
At time zero, some radioactivity was found in the aqueous fraction in experiments with both
anandamide and synaptamide, and these ―blank‖ values limited the sensitivity of the assay. To
determine the extent to which these counts were due to tissue mediated hydrolysis, we
performed experiments using homogenate that had been boiled to destroy enzymatic activity
(fig F.5).
Fig F.4A Fig F.4B Fig F.4C
98
Fig F.5: Comparison of N-acylethanolamine hydrolysis in tissue mediated and tissue independent
environments. T: brain tissue homogenate (20 mg/ml); CT: Coiled brain tissue homogenate (20
mg/ml). Data is expressed in terms of % radiolabeled substrate hydrolyzed and released into the
aqueous layer. N=1.
For both ethanolamides, aqueous counts did increase to a limited extent in incubations using
Coiled tissue. This is presumably due to spontaneous, chemical, hydrolysis under the incubation
conditions. Pretreatment with PF3845 did not reduce aqueous counts in Coiled tissue
incubations to zero, perhaps because of the presence of some [14C]ethanolamine or other
water-soluble impurity in the radiotracer stock solutions. The greater extent of PF-inhibitable
hydrolysis of [14C[synaptamide in unboiled tissue can be attributed to the action of FAAH.
F.3.2.3. Synaptamide inhibits anandamide hydrolysis in crude brain homogenates
Fig F.5A Fig F.5B Fig F.5C
99
We performed a competition binding assay to evaluate whether synaptamide inhibits
anandamide hydrolysis. A tissue concentration of 1.25 mg/ml was chosen to carry out this
experiment as anandamide hydrolysis was linear with time at this concentration. Linearity was
lost at higher concentrations as the substrate was depleted. Incubations were also done with
equimolar mixtures of DHA and ethanolamine to evaluate the extent of possible end-product
inhibition of FAAH.
Competition binding assay
0 1 2 3
0
5
10
15
AEA
EP
DEA
log conc (M)
% s
pe
cif
ic r
ad
iolig
an
d b
ind
ing
Fig F.6: Representative graph showing displacement of [14
C]anandamide binding to FAAH in
mouse brain homogenates (1.25mg/ml; 200 ul) by unlabeled anandamide (AEA), synaptamide
(DEA) or DHA plus ethanolamine (EP). % specific radioligand binding was plotted on y-axis and
log concentrations of the non-radioactive substrates were plotted on x-axis.
Both anandamide and synaptamide inhibited [14C] anandamide hydrolysis, with estimated IC50
values of 8 µM and 68 µM respectively (fig F.6). Mixtures of DHA and ethanolamine had no
effect. This confirms that synaptamide interacts with FAAH but with a lower affinity than
anandamide.
100
F.3.3. End fate of synaptamide
F.3.3.1. Synaptamide incorporates into phospholipids in vivo.
Exogenously administered anandamide in mice is metabolized by FAAH and the released
arachidonic acid is rapidly incorporated into phospholipids (Glaser et al., 2006). We examined
the hydrolysis of exogenous synaptamide and its subsequent incorporation into phospholipids in
mouse brain and blood using 1D-TLC analysis and observed that synaptamide hydrolysis
causes incorporation of the 14C label into phosphatidylethanolmine (fig F.7A and B). This
incorporation was time dependent. Same incorporation pattern was observed with the use of
either [14C-ethanolamine]synaptamide or [14C-docosahexaenoyl]synaptamide.
Fig F.7A Fig F.7B Fig F.7C
101
Fig F.7: Representative 1D-TLC autoradiograph of brain (A) and Blood (B) lipid extract from mice
euthanized after 0, 5, 15, 30 and 60 minutes after the administration of exogenous
[14
C]synaptamide. TLC of pure non-radioactive phospholipids used as a reference for RF values of
various phospholipids relative to each other (C). Mobile phase used:
chloroform:methanol:ammonia 60:30:1. Non-radioactive TLC spots were visualized by exposing
the plate to iodine vapor.
We confirmed that synaptamide ultimately partitions into phosphatidylethanolamine, using 1,2-
dipalmitoyl-sn-glycero-3-phosphoethanolamine (16:0 PE) as our non-radioactive standard
phosphatidylethanolamine. The RF of the PE from brain lipid extract was not the same as our
standard and this is probably due to the difference in fatty acid composition at the sn-1 position
of PE; DHA occupying its sn-2 position (fig F.7).
F.3.3.2. Synaptamide partitions into phospholipids in vitro.
The enzyme FAAH can liberate free fatty acids from fatty acid ethanolamides even in in vitro
systems which are then mainly incorporated into phospholipids (Chicca et al., 2012), (Di Marzo
V, 1994). Results from our experiments also substantiate this observation.
TLC analysis of chloroform extracts from differentiated and undifferentiated N27 cells treated
with [14C-ethanolamine]anandamide and [14C- ethanolamine]synaptamide reveal the chemical
nature of these lipids once taken up into the cell. Comparison with pure radioactive (fatty acid
and fatty acid ethanolamide) and non-radioactive (phospholipid) standards shows that along
102
with corresponding ethanolamides, the radiolabel is incorporated into several other lipids –
phospholipids (fig F.8).
Fig F.8: Representative TLC images of anandamide and synaptamide uptake. Anandamide uptake
without (A) and with PF3845 (B) as well as synaptamide uptake in undifferentiated (A, C and D)
and differentiating (C and E) N27 cells incubated with either 200 nM [14
C-
ethanolamine]anandamide (A,B,C) or [14
C-ethanolamine]synaptamide (D,E).
Since the radiotracers are labeled on the ethanolamine moiety of the respective NAEs, the
ethanolamine released upon their hydrolysis is incorporated into phospholipids –
Fig F.8A Fig F.8B Fig F.8C
Fig F.8D Fig F.8E
103
Phosphatidylethanolamine (PE) and lyso-phosphatidylethanolamine (LYSO-PE) – before it
escapes into the aqueous phase. We confirmed one of the spot corresponds to PE as when
sprayed with ninhydrin, the spot turned a bright pinkish-purple color indicative of a compound
with free amine. In cell lipid extracts along with phosphatidylethanolamine, the hydrolysis
products of synaptamide and anandamide also incorporated into other phospholipids with lower
RF. Since the RFs of phosphatidylcholine, phosphatidylserine and Lyso-
phosphatidylethanolamine were all similar in 1D-TLC, we performed a 2D-TLC to identify which
of these phospholipids incorporates DHA released from synaptamide. Since ninhydrin did not
reveal color in the second phospholipid, we hypothesized it could either be phosphatidylserine
or phosphatidylcholine. Based on the location of the spot on the charred plate loaded with
standard phosphatidylcholine and phosphatidylserine, we confirmed that DHA released from
synaptamide incorporates into PC (fig F.9).
Fig F.9: 2D-TLC of standard non-radioactive phospholipids, PS: phosphatidylserine (A) and PC:
phosphatidylcholine (B) and cell lipid extract of N27 cells incubated with [14
C-EA] Synaptamide
PS PC PC
Fig F.9A Fig F.9 B Fig F.9 C
Dim
ensio
n 1
Dimension 2
104
(C). Mobile phase used: Dimension 1: chloroform:methanol:ammonia 65:35:5; Dimension 2:
chloroform:acetone:methanol:aceticacid:water 30:40:10:10:5. TLC spots are visualized by
charring the plates after spraying with a mixture of copper sulphate and concentrated sulfuric
acid.
Quantification of radioactive TLC spots confirmed a time-dependent increase in intensity in both
undifferentiated and differentiating N27 cells indicating increased incorporation of the label into
the lipid with time; incorporation being higher in undifferentiated cells (fig F.10A). From the TLC
and its quantification, we can confirm that synaptamide partitions into phospholipids, but to a
lesser degree than anandamide consistent with our previous observations that synaptamide
hydrolysis occurs slower than anandamide (fig F.10A).
As free fatty acids are released by FAAH, inhibition of FAAH should decrease the formation of
radiolabeled phospholipids. TLC analysis of lipid extracts of cells pretreated with PF3845 show
that FAAH inhibition significantly decreased the label incorporation into phospholipids and
increased the level of intact ethanolamide in cells (fig F.8 and F.10B).
There is evidence of synaptamide partitioning into phospholipid even when cells were
pretreated with PF3845 suggesting that there is some spontaneous hydrolysis of synaptamide
independent of FAAH (fig F.10B).
105
Fig F.10: Quantification of TLC spots shows the partitioning of anandamide and synaptamide into
phospholipids of cells in the absence (A) or presence (B) of PF3845. Error bars represent SD; N=3
(anandamide), N=4 (synaptamide).
F.3.3.3. Phospholipid partitioning of [14C-ethanolamine]synaptamide versus [14C-
docosahexaenoyl]synaptamide
Fig F.10B
Fig F.10A
106
Using synaptamide or anandamide labeled on the ethanolamine moiety only indicates that the
ethanolamine partitions into phosphatidylethanolamine or lysophosphatidylethanolamine. A
phospholipid can incorporate 2 fatty acids in it – one at sn-1 position (usually a saturated fatty
acid) and another at sn-2 position (usually mono or polyunsaturated fatty acid). bell and brain
extracts inherently have a number of various length chain fatty acids that can incorporate into
the labeled phosphatidylethanolamine. To determine the fate of docosahexaenoyl-chain of
synaptamide, we repeated both in vivo and in vitro experiments with synaptamide labeled on the
docosahexaenoyl moiety of synaptamide.
Fig F.11 A
Fig F.11 B
107
Fig F.11: Quantification of TLC spots shows the partitioning of [14
C-EA] Synaptamide and [14
C-
DHA] Synaptamide into phospholipids of cells in the absence (A) or presence (B) of PF3845. Error
bars represent SD; N=3 ([14
C-DHA] Synaptamide), N=4 ([14
C-EA] Synaptamide).
TLC quantification indicates that in undifferentiated cells both [14C-ethanolamine]synaptamide
and [14C-docosahexaenoyl]synaptamide showed similar incorporation rates into phospholipids;
the rate increasing with time. However, in differentiating cells percentage of phospholipid formed
is higher with [14C-docosahexaenoyl]synaptamide indicating that phospholipid fraction
incorporates the DHA released from synaptamide (fig F.11A). Incubating cells with PF3845 did
inhibit the hydrolysis of synaptamide but not totally (fig F.11B).
F.4. DISCUSSION
The substrate specificity of FAAH is quite selective. The known substrate preference for FAAH
for hydrolysis is amide bond > ester bond, but a mutation at L142 reverses this preference
(ester bond > amide bond) (Patricelli M, 1999). Although FAAH preferentially hydrolyses primary
amides over ethanolamides, it hydrolyses long chain unsaturated fatty acid ethanolamides
faster than saturated fatty acid ethanolamides (Boger D, 2000). The rate of hydrolysis by FAAH
was found to increase with an increase in the degree of unsaturation in the ethanolamide; also,
FAAH preferentially hydrolyses the unsaturated compounds that assume a hair-pin confirmation
(Lang et al., 1999). Anandamide, a good substrate of FAAH, assumes the energy conserving
108
hair pin confirmation, while no such information about synaptamide is known posing a question
as to whether synaptamide is really a substrate for FAAH.
It has been well established that FAAH can hydrolyze a wide range of N-acylethanolamines
such as anandamide and palmitoylethanolamide (Tiger et al., 2000) as well as N-
acylethanolamine analogues (Maccarrone et al., 1998). While evidence supports that
anandamide uptake can be regulated by its metabolism by FAAH (Deutsch et al., 2001), recent
studies show that the activity of FAAH is also influenced by its lipid environment. Anandamide
preferentially associates itself to lipid domains of the ER containing cholesterol. FAAH ―senses‖
the presence of this anandamide localized to the ER and preferentially hydrolyses it and this
preferential selectivity is limited to anandamide (Dainese et al., 2014). The association of
synaptamide with cholesterol is not known yet and this opens up a whole new possibility that the
modest effect of FAAH on synaptamide hydrolysis observed is probably because it is
inaccessible to FAAH. The brain is comprised of a variety of lipid domains consisting of a large
number of possible lipid substrates for FAAH which can bind before synaptamide does, thus
explaining a small role of FAAH in hydrolyzing synaptamide.
Anandamide metabolism into arachidonic acid and ethanolamine was previously demonstrated
in striatal and cortical neurons (Di Marzo V, 1994) as well as in human U937 leukemia cells
(Chicca et al., 2012) among many other cell lines. FAAH expression in these cells has been
documented (Maccarrone et al., 1998; Maccarrone et al., 2004). To date, FAAH expression or
FAAH activity has not been documented in N27 immortalized dopaminergic neural cells. We are
the first to perform a FAAH activity assay in these cells and prove that anandamide is
hydrolyzed in these cells. These cells are particularly interesting because undifferentiated cells
109
are derived from rat fetal mesencephalon and represent ―developing cells‖ and the differentiated
cells undergo morphological changes and represent ―adult neuronal‖ cells. Our uptake results
show a FAAH dependent uptake of anandamide and synaptamide into undifferentiated cells. As
expected, inhibition of FAAH decreased the incorporation of the radiolabel into phospholipids
but this observation is absent in differentiating cells. The uptake of anandamide and
synaptamide into the cell, their metabolism and incorporation into phospholipids is still present,
but all processes seem to be independent of FAAH. The structural changes of N27 cells are still
not known, but it seems like with the onset of differentiation, N27 cells develop a FAAH
independent uptake process probably by inducing the expression of the putative
endocannabinoid transporter.
Many authors use anandamide as a prototype substrate for FAAH and many studies attempted
to establish its Km for FAAH. The km values range from 0.13 μM to 45 μM (Nicolussi et al.,
2015) and this wide variability can be attributed to different assay conditions as well as the fact
that the substrate (anandamide) and final end product (arachidonic acid) are both highly
lipophilic molecules with a tendency to ―stick‖ and form micelles that may modulate the rate of
hydrolysis (Omeir et al., 1995). Thus, in our experiments to identify synaptamide as an inhibitor
of anandamide hydrolysis, we chose tissue and substrate concentrations (1.25 mg/ml and 200
nM respectively) and the incubation time based on the study by Deutsch et al, (Deutsch et al.,
2001) after an exhaustive literature review.
As demonstrated with anandamide, synaptamide hydrolysis releases DHA which is rapidly
incorporated into phospholipids. Consistent with previous studies (Sundler et al., 1974),
exogenous DHA added (either as DHA or by its release from synaptamide) is incorporated into
110
phosphatidylethanolamine and phosphatidylcholine (Nariai et al., 1994). CDP-
ethanolamine:diacylglycerol ethanolamine phosphotransferase, the enzyme involved in the
synthesis of phosphatidylethanolamine preferentially uses diacylglycerols rich in DHA (Kanoh et
al., 1975) explaining the presence of DHA in phosphatidylethanolamine. Since DHA is the
preferred fatty acid for the enzymes for phosphatidylserine (PS) synthesis (Kim et al., 2014), we
expected the cell and brain extracts incubated with synaptamide or DHA would result in the
formation of phosphatidylserine. 2D-TLC of lipid extracts did not reveal the formation of
phosphatidylserine suggesting that longer incubation times are probably required for its
synthesis or that N27 cells lack PSS2 enzymes which preferentially uses
phosphatidylethanolamine with DHA for the synthesis of phosphatidylserine (Kimura et al.,
2013).
F.5. CONCLUSION
Our in vivo uptake studies were consistent with the notion that FAAH can hydrolyze
synaptamide but slowly. While in vivo synaptamide uptake was not significantly impacted by
FAAH inhibition, there is a strong FAAH mediated synaptamide uptake and hydrolysis in N27
cells. This observation is consistent with the studies of Kim et al., who also reported an increase
in the levels of synaptamide in hippocampal neurons after the inhibition of FAAH (Kim et al.,
2011). However, only one study has looked at synaptamide as a potential substrate for FAAH
indirectly by testing its ability to inhibit anandamide hydrolysis in bovine retinas; by competing
with anandamide for FAAH – synaptamide (100 µM) was found to inhibit [14C]anandamide
hydrolysis in bovine retina (Bisogno T, 1999). Our FAAH activity assay results substantiate this
study and reveal that synaptamide at a high concentration is a substrate for FAAH. We further
111
confirmed this by performing a competitive binding assay. Anandamide was potent enough to
bind to the enzyme in 250 µg wet tissue while at least 2 mg of wet tissue was required for FAAH
to have an effect on synaptamide. The IC50 value of synaptamide estimated from our
competition data was about 68 µM, and the physiological level of synaptamide is much less than
that making it an unlikely substrate for FAAH in vivo. This is consistent with the smaller effect of
FAAH inhibition on brain synaptamide uptake in vivo.
112
G. FUNCTIONAL EFFECT OF EXOGENOUS DHA AND SYNAPTAMIDE ON
NEURITOGENESIS AND NEURITE GROWTH
G.1. INTRODUCTION
Neurons are specialized cells with neurites functioning as precursors of axons and dendrites
which play a role in polarization and developing synaptic connections. Neuritogenesis (and
synaptogenesis) occur just prior to the completion of neurogenesis following a cell and a region-
specific agenda (Youdim et al., 2000). Neurite outgrowth is heavily influenced by the
coordination between actin cytoskeleton and microtubular network (Clagett-Dame et al., 2006).
Neurite initiation be gins with the alignment of a dynamic microtubular system towards a point
where actin filaments (in the form of either lamellipodia or filopodia) are present at the periphery
to form a ―growth cone‖. The elongation of the growth cone preferentially occurs in areas where
the actin filopodia surround the actin lamellipodia. This arrangement of actin filaments is
influenced by the interaction between specific ligands (eg. laminin) with their receptors (eg.,
integrins) (da Silva et al., 2002). Actin filaments serve as a guide to direct the microtubular
movement as well as to generate a force within the structure that is stabilized by the
microtubular network (Rodriguez et al., 2003). There are many other signals that can cause
neurite elongation: Activation of AKT and its downstream targets such as glycogen synthase
kinase 3C (GSK3C) the mammalian target of rapamycin (mTOR), cyclic AMP response element
binding protein (CREB), all play a role in neurite outgrowth (Read et al., 2009). Chemokine
CXCL12 interaction with the chemokine receptor CXCR4 is also implicated in axon wiring and
neurite orientation (Yang et al., 2013).
113
The growth of neurites, aside from involving cytoskeletal changes, also involves changes in
surface area. While evidence supports the presence of an inherent elastic stretch that can
increase the surface area of membrane bilayers at physiological conditions, that stretch is not
capable of causing massive changes in surface area that take place during neuritogenesis
(Futerman et al., 1996), suggesting the additional involvement of insertion of new membrane
(Tang, 2001). New membrane insertion corresponds to the synthesis and transport of
membrane proteins and lipids. Membrane proteins are synthesized in rough endoplasmic
reticulum (RER) and the Golgi complex, both of which are localized to the neuronal cell Cody.
Following their synthesis, the membrane proteins are packaged and transported along the
microtubules and possibly along the actin filaments until they reach their site of insertion
(Futerman et al., 1996). The synthesis and transport of membrane lipids is more complex. There
are four main classes of neuronal membrane lipids present in different proportions –
glycerolipids constituting about 60%, sterols constituting about 20%, glycosphingolipids (about
15%) and sphingomyelin (about 5%) (Futerman et al., 1996). Growth cones are rich in smooth
endoplasmic reticulum (SER) (Deitch et al., 1993), which is the main site for synthesis of
glycerophospholipids (mainly phosphatidylcholine) and of cholesterol, indicating that these are
the major lipids synthesized to be incorporated into growing neurites. Using double labeling
experiments with [14C]choline and [3H]choline, Vance and co-workers observed that
approximately 50% of the phospholipid (phosphatidylcholine) was synthesized locally in the
axons (Posse de Chaves et al., 1995). The remaining 50% is probably transported to the axons
from the cell body. Pulse chase experiments with [3H]glycerol confirmed the transport of newly
synthesized glycerophospholipids from the cell body (Pfenninger et al., 1983). The requirement
of the newly synthesized membrane components for insertion into neurites was confirmed when
neurite elongation was blocked on prolonged inhibition of the synthesis of membrane protein
114
(Lein et al., 1991) or lipid (Harel et al., 1993; Posse de Chaves et al., 1995) as well as inhibiting
membrane transport (Osen-Sand et al., 1993).
DHA mediates many of these processes. A study by Green et al. (Green et al., 1999) reported
that just prior to synaptogenesis or neuritogenesis, an increased accumulation of DHA occurred
in rat embryonic neuronal cells. DHA supplementation results in its accumulation into cell body
which is then transported to the nerve growth cones in the form of phospholipids
(phosphatidylethanolamine > phosphatidylserine > phosphatidylcholine > phosphatidylinositol)
for the synthesis of new membrane (Martin, 1998). DHA also induces neuronal differentiation by
decreasing the expression of nestin and promoting the exit of cell cycle (Insua et al., 2003).
Moreover, DHA activates AKT pathway blocking cell death resulting in neurogenesis (Akbar et
al., 2005). DHA promoted neurite elongation initiated by NGF in PC 12 cells (Ikemoto et al.,
1997; Ikemoto et al., 1999). In addition, Calderon and Kim et al. demonstrated that reduced
DHA supplementation to hippocampal primary neuronal cultures resulted in decreased neurite
growth (Calderon et al., 2004). DHA is thought to trigger the activation of many transcriptional
factors by functioning as an endogenous ligand at the retinoid X receptors (RXRs) (de Urquiza
et al., 2000). Upon activation by DHA, RXRs tend to dimerize with other nuclear receptors such
as retinoic acid receptors (RARs), Peroxisome Proliferator-Activated receptors (PPARs) and
Nurr-1 receptors and influence neuronal membrane assembly, synaptic plasticity, cytoskeletal
organization, etc. (Maden, 2002; Wallen-Mackenzie et al., 2003). Combining the evidence in
support of effects of DHA, it can be concluded that it plays a key role in every stage of neuronal
development including neuritogenesis and neurite elongation.
G.2. METHOD
115
G.2.1. Neurite analysis:
We employed a modified version of neurite analysis described by Cao et al. (Cao et al., 2005).
The cover slips with N27 cells mounted on glass slides using DAPI Fluoromount-G were
allowed to dry overnight in a dark place, and then the images for neurite analysis were taken
using an Olympus CX-51 Fluorescent microscope using bright field and DAPI filters.
Approximately 20 fields /cover slip were chosen and only non-clustered differentiated cells were
traced to ensure the precision of the measurements. To minimize bias, all traces were
performed by blinded observers. All raw data that was originally in units of pixels was converted
to units of μm based on a calibration image. A differentiated cell is defined as a cell with at least
one neurite. A neurite refers to any projection from the cell body of the cell which is longer than
the diameter of the cell body from which it projected. Neurite analysis was performed on 70
randomly selected cells and 80 randomly selected neurites from them. Neurite length was
measured using Simple Neurite Tracer plug-in of the NIH imaging software, Image J (version
1.48). The parameters considered for measurement are (1) total number of neurites, (2) total
neurite length per cell, and (3) total length of individual neurites. Neurite number (determined by
counting neurites) and total neurite length per cell (the sum of lengths of all neurites of a cell)
was measured in the randomly selected cells (n=70) with the entire soma and neurites clearly
identifiable. The total length of individual neurites was calculated from the lengths of randomly
selected neurites (n=80) from the 70 differentiated cells. Data from three experiments was
considered for this analysis.
G.3. RESULTS
116
Using an in vitro DHA and synaptamide supplementation approach, we were able to study their
action on neurite growth. N27 cells were cultured for 3 days with differentiating agent and either
DHA or synaptamide at various concentrations. The representative images of undifferentiated
and differentiated cells treated with DAC are shown in Fig. G.1. This enabled us to document
morphological changes that occur in N27 cells during differentiation. Undifferentiated cells are
flat cells with large cytoplasm while differentiated cells have a more polarized morphology with a
large nucleus, little cytoplasm and long neuronal projections: neurites.
Fig G.1: Representative images of undifferentiated (A) and differentiated (C) N27 cells. bells were
stained for tyrosine hydroxylase using rabbit-anti-TH antibody (1:4000; AB192; Chemicon) to
identify the dopaminergic nature of the cell. Tyrosine hydroxylase immunoreactivity was
visualized by using biotinylated goat anti-rabbit secondary IgG antibody (1:250: #BA-1000: Vector
Labs) which after conjugation with Vector ABC reagent (to add avidin-HRP to biotin tag) develops
a brown color on addition of Vector DAB solution.
A B
117
Fig G.2 shows the representative traces of the cell images in image J. These traces were used
to quantify the following parameters - (1) total neurite length, (2) total length of individual
neurites, and (3) total number of neurites.
Fig G.2: The representative images and traces of differentiated N27 cells. (A) Bright field, (B) DAPI,
(C) overlay, (D) traces. The traces were made after calibrating the image scale from pixels to µm.
G.3.1. Effect of synaptamide and DHA on total neurite length in differentiated N27 cells.
The total neurite length (the sum of primary, secondary and tertiary neurites) measured in cells
supplemented with DHA or with synaptamide did not show any significant dose-dependent
changes when compared with control cells; however, different dose-dependent effects occur
between the two. Synaptamide supplementation increased total neurite length at lower
concentrations, (1 nM) but decreased total neurite length at higher concentrations (10 µM).
DHA supplementation, however, showed a slow dose dependent increase in neurite length that
plateaued at higher concentrations (1 nM and 10 µM) (fig G.3). ANOVA analysis followed by
post hoc T-test with Bonferroni‘s correction on the total neurite length of the cells treated with
synaptamide in three different experiments showed that the increased neurite length in cells
TRACES
A B C D
118
treated with low concentrations (1 nM to 100 nM) is significantly higher than in cells treated with
higher concentrations (1 µM and 10 µM). Surprisingly, post hoc t-test with Bonferroni‘s
correction did not show any difference between synaptamide supplemented cells and control
cells (fig G.3).
Fig G.3: Estimation of total neurite length with increasing doses of either DHA or synaptamide.
(N=3; n=70; analysis was done by one-way ANOVA with a post hoc t-test with Bonferroni’s
correction (within synaptamide doses used: *p<0.005 – when compared to total neurite length of
cells treated with 1 nM synaptamide and #p<0.005 when compared to total neurite length of cells
treated with 10 μM synaptamide).
Student‘s T test analysis showed that at lower concentrations (1 nM – 100 nM), the total neurite
length in cells treated with synaptamide is significantly higher than those treated with DHA
(p<0.05) (fig G.4). 10 nM synaptamide seemed to have a maximal effect on neurite elongation.
But since neither lipid showed a significant increase when compared with control cells treated
with just the differentiating agent, this significance becomes questionable.
119
Fig G.4: Comparison of the Total neurite length of N27 cells treated with either DHA or
synaptamide. N=3; n=70; analysis was done by Student’s T test (*p<0.05; comparison between
DHA treated or synaptamide treated cells).
The frequency distribution of total neurite length of the 70 randomly selected cells shows that
lower concentrations of synaptamide (1, 10 or 100 nM) fewer cells tend to have shorter neurites.
In contrast, with higher concentrations of synaptamide (1 µM and 10 µM), the lengths of neurites
are similar to that of control cells. Cells supplemented with DHA have less skew when
compared with synaptamide, even at high concentrations, indicating that their supplementation
did not affect the length of neurites in any way (fig G.5 A and B).
120
Fig G.5: Representative graph of frequency distribution of total neurite length after either DHA (A)
or synaptamide (B) supplementation in differentiated N27 cells. Synaptamide showed a more
skewed neurite distribution with more neurites with a longer length and fewer neurites with
shorter length. (N=1 experiment; n=70 cells).
Total neurite length is the sum of the lengths of all neurites (primary, secondary or tertiary). We
investigated further to see whether the number of neurites per cell, or the lengths of all neurites
were separately altered by supplementation with synaptamide or with DHA.
G.3.2. Effect of synaptamide on individual neurite length of differentiated N27 cells
From the 70 randomly selected cells, 80 neurites were randomly chosen to assess the lengths
of individual neurites. The total lengths were compared across various concentrations of DHA
and synaptamide and we observed the same trend as with total neurite lengths (fig G.6). DHA
treatment increased the length of neurites in a concentration dependent manner but treatment
Fig G.5A Fig G.5 B
121
with synaptamide followed a different trend – there was an initial concentration dependent
increase in neurite length which was lost at higher concentrations
Fig G.6: Estimation of total length of individual neurites selected from randomly selected cells
supplemented with increasing doses of either DHA or synaptamide. (N=3 experiments; n=80
neurites).
We also looked at the frequency distribution of total length of the individual neurites. We found
that supplementation with neither DHA nor synaptamide was associated with the sprouting of
neurites whose lengths did not significantly deviate from those in unsupplemented incubations
(fig G.7A and B).
122
Fig G.7: Representative graph of frequency distribution of individual neurite length after either
DHA (A) or synaptamide (C) supplementation in differentiated N27 cells. (N=1 (representative
image); n=80 neurites).
G.3.3. Effect of synaptamide on the number of individual neurites in differentiated N27 cells
We summed up the number of neurites per cell and plotted a frequency distribution graph to
look at the distribution of the proportion of cells with more neurites. Cells supplemented with
either DHA or synaptamide sprouted similar number of neurites as control cells (fig G.8).
Fig G.7A Fig G.7B
123
Fig G.8: Total number of neurites (sum of primary, secondary and tertiary branches) from N27
cells (n=70) supplemented with either DHA or synaptamide. Data represented as the mean of 3
experiments. Error bars represent SD (N=3 experiments; n=70 cells).
G.3.4. Synaptamide and DHA uptake and metabolism may contribute to their effect on
neuritogenesis and neurite elongation
Based on our neurite analysis, we can conclude that neither synaptamide nor DHA had a
significant effect on neuritogenesis or neurite elongation in N27 cells. We wanted to take a
second look at the functional effect of these lipids on neurite elongation from the perspective of
DHA and synaptamide uptake and metabolism in these cells and try to understand the reason
behind the lack of their effect on neuritogenesis.
Radiolabeled synaptamide and DHA were added to N27 cells at a concentration of 200 nM in
order to study their uptake. Hence, the effect of these lipids at a concentration of 100 nM was
124
closely re-examined to explain the observed effects on neuritogenesis and neurite elongation.
At 100 nM, the total neurite length of the cells, the lengths of individual neurites as well as the
number of neurites with synaptamide and DHA were similar. There was no statistical
significance between the two treatments (fig G.9).
Fig G.9: Comparison of neurite lengths (A) and the number of neurites (C) from randomly selected
N27 cells (n=70) treated with either DHA 100 nM or synaptamide 100 nM. Data represented as the
mean of 3 experiments. Error bars represent SD (N=3 experiments; n=70 cells).
This observation was consistent with our uptake studies. DHA and synaptamide are taken into
the cells at similar rates but with the onset of differentiation, DHA is taken up preferentially over
synaptamide. In both undifferentiated and differentiating cells, DHA and synaptamide partition
into phospholipids to the same extent (fig G.10) suggesting that both contribute to the synthesis
of membrane phospholipids to the same extent. This may explain, at least in part, the similar
effect of both DHA and synaptamide on neurite elongation in N27 cells. Although the
Fig G.9A Fig G.9B
125
incorporation of DHA into phospholipids was higher than for synaptamide at early times, the
advantage was lost at later time-points (fig G.10).
Fig G.10: Time dependent phospholipid incorporation of exogenous [14
C]DHA and [14
C-
docosahexaenoyl]synaptamide in undifferentiated (A) and differentiating (C) N27 cells. Data
represented as the average % of radioactivity incorporated into phospholipids as quantified from
autoradiographs. Error bars represent SD (N=3 experiments; n=70 cells).
G.4. DISCUSSION
Synaptamide was found to play a role in neuritogenesis and neurite elongation in hippocampal
cultures (Kim et al., 2011). Based on the results from studies in hippocampal neurons, we
evaluated whether synaptamide and/or DHA increased neurite growth in N27 cells. N27 cells
are dopaminergic immortalized mesencephalic neural cells (Clarkson et al., 1999). Our data
indicate that neurite elongation was not increased by either synaptamide or by DHA.
Fig G.10A Fig G.10B
126
The doses of synaptamide and DHA used for supplementing the cell culture were based on
previous studies which demonstrated a functional effect (Kim et al., 2011). We employed a dose
range of 1 nM to 10 µM of DHA and synaptamide for our analysis and even at concentrations as
high as 10 µM the effect of DHA was similar to that of control cells. This observation was in
contrast to previous studies that reported a toxic effect of DHA at 5 µM concentration on
hippocampal neurons (Cao et al., 2009). This discrepancy may be due to the fact that we used
an immortalized cell line instead of a primary culture. PUFAs are known to generate free
radicals which can result in cell death in primary neuron cultures. The N27 cells were treated
with dibutyryl-cAMP and dehydroepiandrosterone (a precursor for androgens) prior to adding
DHA or synaptamide. Androgen pretreatment in N27 cells is known to condition them against
oxidative damage (Holmes et al., 2013) and addition of dehydroepiandrosterone may possibly
prevent these cells from toxic effects of high concentrations of DHA and synaptamide. In
addition to this, we also supplemented our cultures with 40 µM α-tocopherol to minimize the
oxidation of DHA and synaptamide.
The lack of effect of DHA and synaptamide on neurite growth may be explained on the basis of
their neuronal uptake and metabolism. At 200 nM, both DHA and synaptamide were rapidly
taken up by the N27 cells and were incorporated into phospholipids. The incorporation of DHA
into phospholipids occurs to the same extent from either free acid or synaptamide. If DHA and
synaptamide contribute to new membrane synthesis, neurite growth will occur to the same
extent. The slight increase in neurite elongation by synaptamide may suggest an independent
action by functioning as a signaling molecule, but the lack of significance when compared with
the control cells rule out this possibility.
127
Since DHA also did not cause neurite elongation in N27 cells, it can be suggested that N27 cells
have a different mechanism by which neurite elongation may take place. N27 cells are
immortalized rat fetal mesencephalic cells and even though they express neuron specific
markers, they are not considered as neurons. The differences between immortalized cells and
primary neuron cultures, while can be useful in some studies, may not be the best models for
determining neurogenic potential of growth factors.
G.5. CONCLUSION
Overall, there were no significant differences between synaptamide supplemented or control
(unsupplemented) cells or between DHA supplemented and control cells in terms of total neurite
length, number of neurites per cell, or lengths of individual neurites. In vitro systems provide a
great alternative platform to understand mechanisms underlying physiological/ pathological
processes in a simplified system which, in vivo are much more complex. The use of in vitro
neuronal systems facilitated the understanding the functioning of nervous system which is
challenging because mature neurons do not undergo cell division. The use of primary and
secondary neuronal cultures made advancement possible in understanding the functioning of
nervous systems (Gordon et al., 2013). Most primary cultures are derived from embryonic
tissues with limited cell-division properties. They are useful as they ―mimic‖ the properties of
neurons in vivo, but cannot be used in all studies due to the disadvantage of being difficult to
manipulate and study the effect of therapeutic interventions (Aldrich, 2016). Secondary neuronal
cultures are more useful as they can be manipulated easily to test various end points.
Secondary cultures are derived from fetal cells or neuronal tumors and are manipulated to
become immortalized so they can be grown easily with minimal variability between passages.
128
As these cultures are derived from variant sources, their properties differ considerably from
neuronal cells in vivo (Welser-Alves, 2015). While addition of growth factors and other
transcription factors can induce a more ―neuronal‖ phenotype, they can not necessarily be used
to recapitulate the effect of these factors as observed in vivo. Our study was consistent with this
observation. The effects seen in vitro were only partially translatable to in vivo conditions.
129
H. CONCLUDING REMARKS
The overall goal of this dissertation project was to examine the metabolism and biological
effects of N-docosahexaenoylethanolamine (DHA-ethanolamine; ―synaptamide‖). This
compound is structurally similar to the endocannabinoid, anandamide (N-
arachidonoylethanolamine), but incorporates the omega-3 polyunsaturated fatty acid,
docosahexaenoic acid (DHA) in place of the omega-6 fatty acid, arachidonic acid. Unlike
anandamide, synaptamide has very low affinity for cannabinoid receptors, but has recently been
proposed to be important for synapse-formation. Both arachidonic acid and DHA are major
contributors to the brain‘s content of esterified long chain fatty acid, and are therefore important
structural components of brain membranes. Arachidonic acid is the precursor of lipid signaling
molecules such as prostaglandins that are produced via the action of oxygenases, in addition to
its role as a precursor of endocannabinoids. Lipid signaling molecules derived from DHA are
less well understood than those derived from arachidonic acid.
The dissertation is divided into six sections, A—F.
Section A presents the biological background to the project. An important starting point was the
suggestion by Kim et al. (2011) that some effects of exogenous DHA might be mediated via
production of synaptamide.
Section C presents the experimental methods used. These included: administration of
radiolabeled synaptamide to mice, with and without pharmacological pre-treatment; brain
microdissection studies; tissue culture experiments using undifferentiated and differentiated N27
130
neural cells; tissue extraction and radiochromatography of radiolabeled lipids produced from
labeled synaptamide and DHA; enzyme kinetic studies; and studies of uptake of labeled DHA
and synaptamide by cells.
In Section C, studies are described in which radiochromatographic evidence was sought for
biosynthesis of synaptamide from DHA. Analogous studies were also done to search for
production of anandamide from arachidonic acid. Following intravenous injection of carbon-14
labeled arachidonic acid and DHA to mice, radiolabeled anandamide and synaptamide,
respectively were detected in brain extracts. However, in incubations with N27 cells, labeled
anandamide was produced from labeled arachidonic acid, but labeled synaptamide was not
found in extracts of cells that had been incubated with labeled DHA. Thus it appears that the
suggestion of the Kim group that effects of DHA may be mediated via synaptamide formation is
not true, at least in N27 cells.
Section D describes in vivo and tissue culture experiments in which uptake of synaptamide,
labeled either in the DHA moiety or the ethanolamine moiety, and the formation of labeled DHA
and ethanolamine from these radiotracers, was examined. Labeled DHA and ethanolamine
were also employed in uptake experiments. Brain uptake of labeled synaptamide, expressed as
percent injected radioactivity per gram of tissue, was found to be greater for synaptamide than
for DHA, in agreement with previous studies of labeled anandamide and arachidonic acid in the
Gatley laboratory.
The main subject of Section E is metabolism of synaptamide by the enzyme fatty acid amide
hydrolase (FAAH). This enzyme terminates action of the endocannabinoid, anandamide, by
131
cleaving its amide bond. In studies with mouse brain homogenates, it was found that
synaptamide was a poorer inhibitor of radioactive anandamide hydrolysis than was anandamide
itself. IC50 values were a factor-of-ten higher for synaptamide than anandamide. Using
radiolabeled synaptamide, enzyme-mediated hydrolysis was documented, but at a slower rate
than for anandamide. In vivo, radiolabeled phospholipids were found in brain after
administration of labeled synaptamide, confirming hydrolysis of synaptamide in vivo.
Pretreatment of mice with a potent inhibitor of FAAH significantly reduced but did not totally
eliminate formation of labeled phospholipids, suggesting some non-FAAH mediated hydrolysis
of synaptamide.
In Section F, tissue culture experiments in the N27 cells are describe d in which effects of
synaptamide and DHA on cell morphology were compared. There were no significant
differences between synaptamide supplemented or control (unsupplemented) cells or between
DHA supplemented and control cells in terms of total neurite length, number of neurites per cell,
or lengths of individual neurites. However, there were apparent significant differences (p
<0.005) between cells supplemented at 10 nM or 100 nM with DHA or with synaptamide for total
neurite length. One could speculate that DHA and synaptamide have opposing effects on
neuritogenesis at these concentrations, but in retrospect it is clear that N27 cells are not a good
model to evaluate Kim‘s inference (based on primary hippocampal cultures) that DHA acts via
synaptamide, since, as noted in Section C, we did not find [14C]synaptamide in N27 cells
incubated with [14C]DHA.
Much remains unclear about synaptamide that should be investigated in further studies. Thus
far, one group (Kim and co-workers) has been the main contributor in investigating the possible
132
therapeutic potential of synaptamide. The findings in this dissertation are consistent with the
assumption that synaptamide has a similar biosynthetic pathway to that of anandamide, but do
not directly clarify the issue of whether documented effects of DHA in vivo are mediated via
synaptamide.
133
I. REFERENCES:
1. Aarnisalo P, Kim CH, Lee JW, Perlmann T (2002). Defining requirements for heterodimerization between the retinoid X receptor and the orphan nuclear receptor Nurr1. The Journal of biological chemistry 277(38): 35118-35123.
2. Abumrad N, Coburn C, Ibrahimi A (1999). Membrane proteins implicated in long-chain
fatty acid uptake by mammalian cells: CD36, FATP and FABPm. Biochimica et biophysica acta 1441(1): 4-13.
3. Adams FS, La Rosa FG, Kumar S, Edwards-Prasad J, Kentroti S, Vernadakis A, et al.
(1996). Characterization and transplantation of two neuronal cell lines with dopaminergic properties. Neurochemical research 21(5): 619-627.
4. Adibhatla RM, Hatcher JF (2007). Role of Lipids in Brain Injury and Diseases. Future
lipidology 2(4): 403-422.
5. Akbar M, Calderon F, Wen Z, Kim HY (2005). Docosahexaenoic acid: a positive modulator
of Akt signaling in neuronal survival. Proceedings of the National Academy of Sciences of the United States of America 102(31): 10858-10863.
6. Aldrich S (2016). Primary cell Culture Vol. 2016.
7. Balvers MG, Verhoeckx KC, Plastina P, Wortelboer HM, Meijerink J, Witkamp RF (2010).
Docosahexaenoic acid and eicosapentaenoic acid are converted by 3T3-L1 adipocytes to N-acyl ethanolamines with anti-inflammatory properties. Biochim Biophys Acta 1801(10): 1107-1114.
8. Banker AHFaGA (1996). The economics of neurite outgrowth - the addition of new
membrane to growing axons. Trends Neurosci.(19): 144-149.
9. Beltramo M, Stella N, Calignano A, Lin SY, Makriyannis A, Piomelli D (1997). Functional
role of high-affinity anandamide transport, as revealed by selective inhibition. Science 277(5329): 1094-1097.
134
10. Bisogno T D-VI, Lagarde M, Di Marzo V. (1999). Biosynthesis and Inactivation of N-Arachidonoylethanolamine (Anandamide) and N-Docosahexaenoylethanolamine in Bovine Retina. Archives of Biochemistry and Biophysics 370(2): 300-307.
11. Boger D FR, Patterson J, Cravatt B. (2000). Fatty acid amide hydrolase substrate
specificity. bioorganic and medicinal chemistry letters 10: 2613-2616.
12. Bojesen IN, Hansen HS (2003). Binding of anandamide to bovine serum albumin. Journal
of lipid research 44(9): 1790-1794.
13. Brown I, Cascio MG, Wahle KW, Smoum R, Mechoulam R, Ross RA, et al. (2010).
Cannabinoid receptor-dependent and -independent anti-proliferative effects of omega-3 ethanolamides in androgen receptor-positive and -negative prostate cancer cell lines. Carcinogenesis 31(9): 1584-1591.
14. Calderon F, Kim HY (2004). Docosahexaenoic acid promotes neurite growth in
hippocampal neurons. J Neurochem 90(4): 979-988.
15. Cao D, Kevala K, Kim J, Moon HS, Jun SB, Lovinger D, et al. (2009). Docosahexaenoic acid
promotes hippocampal neuronal development and synaptic function. J Neurochem 111(2): 510-521.
16. Cao D, Xue R, Xu J, Liu Z (2005). Effects of docosahexaenoic acid on the survival and
neurite outgrowth of rat cortical neurons in primary cultures. The Journal of nutritional biochemistry 16(9): 538-546.
17. Chang MC, Arai T, Freed LM, Wakabayashi S, Channing MA, Dunn BB, et al. (1997). Brain
incorporation of [1-11C]arachidonate in normocapnic and hypercapnic monkeys, measured with positron emission tomography. Brain Res 755(1): 74-83.
18. Chicca A, Marazzi J, Nicolussi S, Gertsch J (2012). Evidence for bidirectional
endocannabinoid transport across cell membranes. The Journal of biological chemistry 287(41): 34660-34682.
19. Christie WW An Introduction to Lipids and Gas Chromatography - AOCS Lipid Library:
CHAP 1. AOCS Lipid Library.
135
20. Clagett-Dame M, McNeill EM, Muley PD (2006). Role of all-trans retinoic acid in neurite outgrowth and axonal elongation. Journal of neurobiology 66(7): 739-756.
21. Clandinin MT, Chappell JE, Leong S, Heim T, Swyer PR, Chance GW (1980). Intrauterine
fatty acid accretion rates in human brain: implications for fatty acid requirements. Early human development 4(2): 121-129.
22. Clarkson ED, Edwards-Prasad J, Freed CR, Prasad KN (1999). Immortalized dopamine
neurons: A model to study neurotoxicity and neuroprotection. Proceedings of the Society for Experimental Biology and Medicine. Society for Experimental Biology and Medicine 222(2): 157-163.
23. Contreras MA, Greiner RS, Chang MC, Myers CS, Salem N, Jr., Rapoport SI (2000).
Nutritional deprivation of alpha-linolenic acid decreases but does not abolish turnover and availability of unacylated docosahexaenoic acid and docosahexaenoyl-CoA in rat brain. J Neurochem 75(6): 2392-2400.
24. Cravatt BF, Giang DK, Mayfield SP, Boger DL, Lerner RA, Gilula NB (1996). Molecular
characterization of an enzyme that degrades neuromodulatory fatty-acid amides. Nature 384(6604): 83-87.
25. Croset B, Polette, Lagarde (2000). Characterization of plasma unsaturated
lysophosphatidylcholines in human and rat. Biochem. J. 345: 61–67.
26. da Silva JS, Dotti CG (2002). Breaking the neuronal sphere: regulation of the actin
cytoskeleton in neuritogenesis. Nature reviews. Neuroscience 3(9): 694-704.
27. Dainese E, De Fabritiis G, Sabatucci A, Oddi S, Angelucci CB, Di Pancrazio C, et al. (2014).
Membrane lipids are key modulators of the endocannabinoid-hydrolase FAAH. The Biochemical journal 457(3): 463-472.
28. Day TA, Rakhshan F, Deutsch DG, Barker EL (2001). Role of fatty acid amide hydrolase in
the transport of the endogenous cannabinoid anandamide. Molecular pharmacology 59(6): 1369-1375.
29. de Urquiza AM, Liu S, Sjoberg M, Zetterstrom RH, Griffiths W, Sjovall J, et al. (2000).
Docosahexaenoic acid, a ligand for the retinoid X receptor in mouse brain. Science 290(5499): 2140-2144.
136
30. Deitch JS, Banker GA (1993). An electron microscopic analysis of hippocampal neurons
developing in culture: early stages in the emergence of polarity. The Journal of neuroscience : the official journal of the Society for Neuroscience 13(10): 4301-4315.
31. Deutsch DG (2016). A Personal Retrospective: Elevating Anandamide (AEA) by Targeting
Fatty Acid Amide Hydrolase (FAAH) and the Fatty Acid Binding Proteins (FABPs). Frontiers in pharmacology 7: 370.
32. Deutsch DG, Chin SA (1993). Enzymatic synthesis and degradation of anandamide, a
cannabinoid receptor agonist. Biochem Pharmacol 46(5): 791-796.
33. Deutsch DG, Glaser ST, Howell JM, Kunz JS, Puffenbarger RA, Hillard CJ, et al. (2001). The
cellular uptake of anandamide is coupled to its breakdown by fatty-acid amide hydrolase. The Journal of biological chemistry 276(10): 6967-6973.
34. Devane W A AJ (1994). Enzymatic synthesis of anandamide, an endogenous ligand for
the cannabinoid receptor, by brain membranes. Proc. Natl. Acad. Sci 91: 6698-6701.
35. Di Marzo V FA, Piomelli D (1994). Formation and inactivation of endogenous
cannabinoid anandamide in central neurons. Nature 372: 686-691.
36. Diana MA, Marty A (2004). Endocannabinoid-mediated short-term synaptic plasticity:
depolarization-induced suppression of inhibition (DSI) and depolarization-induced suppression of excitation (DSE). British journal of pharmacology 142(1): 9-19.
37. Farooqui AA, Horrocks LA (2004). Brain phospholipases A2: a perspective on the history.
Prostaglandins, leukotrienes, and essential fatty acids 71(3): 161-169.
38. Fasia L, Karava V, Siafaka-Kapadai A (2003). Uptake and metabolism of [3H]anandamide
by rabbit platelets. European Journal of Biochemistry 270(17): 3498-3506.
39. Felder CC, Briley EM, Axelrod J, Simpson JT, Mackie K, Devane WA (1993). Anandamide,
an endogenous cannabimimetic eicosanoid, binds to the cloned human cannabinoid receptor and stimulates receptor-mediated signal transduction. Proceedings of the National Academy of Sciences of the United States of America 90(16): 7656-7660.
137
40. Fowler CJ (2012). Anandamide uptake explained? Trends in pharmacological sciences 33(4): 181-185.
41. Fowler CJ, Tiger G, Ligresti A, Lopez-Rodriguez ML, Di Marzo V (2004). Selective
inhibition of anandamide cellular uptake versus enzymatic hydrolysis--a difficult issue to handle. European journal of pharmacology 492(1): 1-11.
42. Freund T F KI, Piomelli D (2003). Role of Endogenous Cannabinoids in Synaptic Signaling.
Physiol Rev 83: 1017–1066.
43. Futerman AH, Banker GA (1996). The economics of neurite outgrowth--the addition of
new membrane to growing axons. Trends in neurosciences 19(4): 144-149.
44. Galligan JJ (2009). Cannabinoid signalling in the enteric nervous system.
Neurogastroenterology and motility : the official journal of the European Gastrointestinal Motility Society 21(9): 899-902.
45. Giuffrida A, Parsons LH, Kerr TM, Rodriguez de Fonseca F, Navarro M, Piomelli D (1999).
Dopamine activation of endogenous cannabinoid signaling in dorsal striatum. Nature neuroscience 2(4): 358-363.
46. Giuffrida A, Rodriguez de Fonseca F, Nava F, Loubet-Lescoulie P, Piomelli D (2000).
Elevated circulating levels of anandamide after administration of the transport inhibitor, AM404. European journal of pharmacology 408(2): 161-168.
47. Glaser ST, Gatley SJ, Gifford AN (2006). Ex vivo imaging of fatty acid amide hydrolase
activity and its inhibition in the mouse brain. J Pharmacol Exp Ther 316(3): 1088-1097.
48. Glaser ST, Kaczocha M, Deutsch DG (2005). Anandamide transport: a critical review. Life
sciences 77(14): 1584-1604.
49. Goparaju SK, Ueda N, Yamaguchi H, Yamamoto S (1998). Anandamide amidohydrolase
reacting with 2-arachidonoylglycerol, another cannabinoid receptor ligand. FEBS Lett 422(1): 69-73.
50. Gordon J, Amini S, White MK (2013). General overview of neuronal cell culture. Methods
in molecular biology 1078: 1-8.
138
51. Green P, Glozman S, Kamensky B, Yavin E (1999). Developmental changes in rat brain
membrane lipids and fatty acids. The preferential prenatal accumulation of docosahexaenoic acid. Journal of lipid research 40(5): 960-966.
52. Green P, Yavin E (1996). Fatty acid composition of late embryonic and early postnatal rat
brain. Lipids 31(8): 859-865.
53. Gulyas AI, Cravatt BF, Bracey MH, Dinh TP, Piomelli D, Boscia F, et al. (2004). Segregation
of two endocannabinoid-hydrolyzing enzymes into pre- and postsynaptic compartments in the rat hippocampus, cerebellum and amygdala. The European journal of neuroscience 20(2): 441-458.
54. Hamilton JA, Johnson RA, Corkey B, Kamp F (2001). Fatty acid transport: the diffusion
mechanism in model and biological membranes. Journal of molecular neuroscience : MN 16(2-3): 99-108; discussion 151-107.
55. Harel R, Futerman AH (1993). Inhibition of sphingolipid synthesis affects axonal
outgrowth in cultured hippocampal neurons. The Journal of biological chemistry 268(19): 14476-14481.
56. Hillard C J EWS, Campbell W B. (1997). Accumulation of N-Arachidonoylethanolamine
(Anandamide) into be rebellar Granule cells Occurs via Facilitated DIffusion. Journal of Neurochemistry 69: 631-638.
57. Hillard CJ, Jarrahian A (2003). be llular accumulation of anandamide: consensus and
controversy. British journal of pharmacology 140(5): 802-808.
58. Holmes S, Abbassi B, Su C, Singh M, Cunningham RL (2013). Oxidative stress defines the
neuroprotective or neurotoxic properties of androgens in immortalized female rat dopaminergic neuronal cells. Endocrinology 154(11): 4281-4292.
59. Hong S, Gronert K, Devchand PR, Moussignac RL, Serhan CN (2003). Novel docosatrienes
and 17S-resolvins generated from docosahexaenoic acid in murine brain, human blood, and glial cells. Autacoids in anti-inflammation. The Journal of biological chemistry 278(17): 14677-14687.
139
60. Hu K EXPLAINING THE PATTERNS OF DISTRIBUTION OF EXOGENOUS RADIOLABELED ANANDAMIDE AND RELATED COMPOUNDS IN THE MOUSE BRAIN. PhD Doctoral Dissertation, Northeastern Unversity, Boston, MA, 2016.
61. Hu K, Sonti S, Glaser ST, Duclos RI, Jr., Gatley SJ (In press). Brain Uptake And Metabolism
Of The Endocannabinoid Anandamide Labeled In Either The Arachidonoyl Or Ethanolamine Moiety. Nuclear Medicine and Biology.
62. Huerlimann R, Steinig EJ, Loxton H, Zenger KR, Jerry DR, Heimann K (2014). Effects of
growth phase and nitrogen starvation on expression of fatty acid desaturases and fatty acid composition of Isochrysis aff. galbana (TISO). Gene 545(1): 36-44.
63. Ikemoto A, Kobayashi T, Emoto K, Umeda M, Watanabe S, Okuyama H (1999). Effects of
docosahexaenoic and arachidonic acids on the synthesis and distribution of aminophospholipids during neuronal differentiation of PC12 cells. Arch Biochem Biophys 364(1): 67-74.
64. Ikemoto A, Kobayashi T, Watanabe S, Okuyama H (1997). Membrane fatty acid
modifications of PC12 cells by arachidonate or docosahexaenoate affect neurite outgrowth but not norepinephrine release. Neurochemical research 22(6): 671-678.
65. Insua MF, Garelli A, Rotstein NP, German OL, Arias A, Politi LE (2003). cell bycle
regulation in retinal progenitors by glia-derived neurotrophic factor and docosahexaenoic acid. Investigative ophthalmology & visual science 44(5): 2235-2244.
66. Jacobsson SO, Fowler CJ (2001). Characterization of palmitoylethanolamide transport in
mouse Neuro-2a neuroblastoma and rat RBL-2H3 basophilic leukaemia cells: comparison with anandamide. British journal of pharmacology 132(8): 1743-1754.
67. Jin XH, Okamoto Y, Morishita J, Tsuboi K, Tonai T, Ueda N (2007). Discovery and
characterization of a Ca2+-independent phosphatidylethanolamine N-acyltransferase generating the anandamide precursor and its congeners. The Journal of biological chemistry 282(6): 3614-3623.
68. Kalant D CK (2004). Regulation of fatty acid transport. Curr Opin Lipidol 15: 309-314.
140
69. Kamp F, Hamilton JA (2006). How fatty acids of different chain length enter and leave cells by free diffusion. Prostaglandins, leukotrienes, and essential fatty acids 75(3): 149-159.
70. Kan I, Melamed E, Offen D, Green P (2007). Docosahexaenoic acid and arachidonic acid
are fundamental supplements for the induction of neuronal differentiation. Journal of lipid research 48(3): 513-517.
71. Kang MJ, Fujino T, Sasano H, Minekura H, Yabuki N, Nagura H, et al. (1997). A novel
arachidonate-preferring acyl-CoA synthetase is present in steroidogenic cells of the rat adrenal, ovary, and testis. Proceedings of the National Academy of Sciences of the United States of America 94(7): 2880-2884.
72. Kanoh H, Ohno K (1975). Substrate-selectivity of rat liver microsomal 1,2-diacylglycerol:
CDP-choline(ethanolamine) choline(ethanolamine)phosphotransferase in utilizing endogenous substrates. Biochimica et biophysica acta 380(2): 199-207.
73. Kathuria S, Gaetani S, Fegley D, Valino F, Duranti A, Tontini A, et al. (2003). Modulation
of anxiety through blockade of anandamide hydrolysis. Nature medicine 9(1): 76-81.
74. Kim HY (2007). Novel metabolism of docosahexaenoic acid in neural cells. The Journal of
biological chemistry 282(26): 18661-18665.
75. Kim HY, Bigelow J, Kevala JH (2004). Substrate preference in phosphatidylserine
biosynthesis for docosahexaenoic acid containing species. Biochemistry 43(4): 1030-1036.
76. Kim HY, Huang BX, Spector AA (2014). Phosphatidylserine in the brain: metabolism and
function. Progress in lipid research 56: 1-18.
77. Kim HY, Moon HS, Cao D, Lee J, Kevala K, Jun SB, et al. (2011). N-
Docosahexaenoylethanolamide promotes development of hippocampal neurons. The Biochemical journal 435(2): 327-336.
78. Kimura AK, Kim HY (2013). Phosphatidylserine synthase 2: high efficienby for
synthesizing phosphatidylserine containing docosahexaenoic acid. Journal of lipid research 54(1): 214-222.
141
79. Kruszka KK, Gross RW (1994). The ATP- and CoA-independent synthesis of
arachidonoylethanolamide. A novel mechanism underlying the synthesis of the endogenous ligand of the cannabinoid receptor. The Journal of biological chemistry 269(20): 14345-14348.
80. Lagarde M BN, Brossard M, Croset M, Lecerf J. (2000). Lysophosphatidylcholine as a
Preferred Carrier Form of Docosahexaenoic Acid to the Brain. Journal of Molecular Neuroscience 1(16): 201–204.
81. Lamaziere A, Richard D, Barbe U, Kefi K, Bausero P, Wolf C, et al. (2011). Differential
distribution of DHA-phospholipids in rat brain after feeding: A lipidomic approach. Prostaglandins, leukotrienes, and essential fatty acids 84(1-2): 7-11.
82. Lane MA, Bailey SJ (2005). Role of retinoid signalling in the adult brain. Progress in
neurobiology 75(4): 275-293.
83. Lang W, Qin C, Lin S, Khanolkar AD, Goutopoulos A, Fan P, et al. (1999). Substrate
specificity and stereoselectivity of rat brain microsomal anandamide amidohydrolase. Journal of medicinal chemistry 42(5): 896-902.
84. Lein PJ, Higgins D (1991). Protein synthesis is required for the initiation of dendritic
growth in embryonic rat sympathetic neurons in vitro. Brain research. Developmental brain research 60(2): 187-196.
85. Leung K, Elmes MW, Glaser ST, Deutsch DG, Kaczocha M (2013). Role of FAAH-like
anandamide transporter in anandamide inactivation. PloS one 8(11): e79355.
86. Liu J, Wang L, Harvey-White J, Huang BX, Kim HY, Luquet S, et al. (2008). Multiple
pathways involved in the biosynthesis of anandamide. Neuropharmacology 54(1): 1-7.
87. Lo Van A, Sakayori N, Hachem M, Belkouch M, Picq M, Lagarde M, et al. (2016).
Mechanisms of DHA transport to the brain and potential therapy to neurodegenerative diseases. Biochimie.
88. Lukiw WJ, Cui JG, Marcheselli VL, Bodker M, Botkjaer A, Gotlinger K, et al. (2005). A role
for docosahexaenoic acid-derived neuroprotectin D1 in neural cell survival and Alzheimer disease. The Journal of clinical investigation 115(10): 2774-2783.
142
89. Maccarrone M, Gasperi V, Fezza F, Finazzi-Agro A, Rossi A (2004). Differential regulation
of fatty acid amide hydrolase promoter in human immune cells and neuronal cells by leptin and progesterone. European journal of biochemistry / FEBS 271(23-24): 4666-4676.
90. Maccarrone M, van der Stelt M, Rossi A, Veldink GA, Vliegenthart JF, Agro AF (1998).
Anandamide hydrolysis by human cells in culture and brain. The Journal of biological chemistry 273(48): 32332-32339.
91. Maden M (2002). Retinoid signalling in the development of the central nervous system.
Nature reviews. Neuroscience 3(11): 843-853.
92. Marszalek JR, Kitidis C, Dirusso CC, Lodish HF (2005a). Long-chain acyl-CoA synthetase 6
preferentially promotes DHA metabolism. The Journal of biological chemistry 280(11): 10817-10826.
93. Marszalek JR, Lodish HF (2005b). Docosahexaenoic acid, fatty acid-interacting proteins,
and neuronal function: breastmilk and fish are good for you. Annual review of cell and developmental biology 21: 633-657.
94. Martin RE (1998). Docosahexaenoic acid decreases phospholipase A2 activity in the
neurites/nerve growth cones of PC12 cells. J Neurosci Res 54(6): 805-813.
95. Martinez M, Mougan I (1998). Fatty acid composition of human brain phospholipids
during normal development. J Neurochem 71(6): 2528-2533.
96. Mashek DG, Li LO, Coleman RA (2007). Long-chain acyl-CoA synthetases and fatty acid
channeling. Future lipidology 2(4): 465-476.
97. Maurelli s BT, Di Marzo V. (1995). Two novel classes of neuroactive fatty acid amides are
substrates for mouse neuroblastoma 'anandamide amidohydrolase'. FEBS Letters 377: 82-86.
98. McAndrew RS, Chapman KD (1998). Enzymology of cottonseed microsomal N-
acylphosphatidylethanolamine synthase: kinetic properties and mechanism-based inactivation. Biochimica et biophysica acta 1390(1): 21-36.
143
99. Mechoulam R, Fride E, Di Marzo V (1998). Endocannabinoids. European journal of
pharmacology 359(1): 1-18.
100. Montuschi P, Barnes PJ, Roberts LJ, 2nd (2004). Isoprostanes: markers and mediators of
oxidative stress. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 18(15): 1791-1800.
101. Moore SA, Yoder E, Murphy S, Dutton GR, Spector AA (1991). Astrobytes, not neurons,
produce docosahexaenoic acid (22:6 omega-3) and arachidonic acid (20:4 omega-6). Journal of neurochemistry 56(2): 518-524.
102. Morash S C CHW, Spence M W (1989). Lysophosphatidylcholine as an intermediate in
phosphatidylcholine metabolism and glycerophosphocholine synthesis in cultured cells: an evaluation of the roles of 1-acyl- and 2-acyl-lysophosphatidylcholine. Biochim Biophys Acta(1004): 221-229.
103. Muller CP, Reichel M, Muhle C, Rhein C, Gulbins E, Kornhuber J (2015). Brain membrane
lipids in major depression and anxiety disorders. Biochimica et biophysica acta 1851(8): 1052-1065.
104. Nakamura MT, Nara TY (2004). Structure, function, and dietary regulation of delta6,
delta5, and delta9 desaturases. Annual review of nutrition 24: 345-376.
105. Nariai T, DeGeorge JJ, Greig NH, Genka S, Rapoport SI, Purdon AD (1994). Differences in
rates of incorporation of intravenously injected radiolabeled fatty acids into phospholipids of intracerebrally implanted tumor and brain in awake rats. Clinical & experimental metastasis 12(3): 213-225.
106. Nguyen LN, Ma D, Shui G, Wong P, Cazenave-Gassiot A, Zhang X, et al. (2014). Mfsd2a is
a transporter for the essential omega-3 fatty acid docosahexaenoic acid. Nature 509(7501): 503-506.
107. Nicolussi S, Gertsch J (2015). Endocannabinoid transport revisited. Vitamins and
hormones 98: 441-485.
108. Oddi S, Bari M, Battista N, Barsacchi D, Cozzani I, Maccarrone M (2005). Confocal
microscopy and biochemical analysis reveal spatial and functional separation between
144
anandamide uptake and hydrolysis in human keratinobytes. be llular and molecular life sciences : CMLS 62(3): 386-395.
109. Oddi S, Fezza F, Catanzaro G, De Simone C, Pucci M, Piomelli D, et al. (2010). Pitfalls and
solutions in assaying anandamide transport in cells. Journal of lipid research 51(8): 2435-2444.
110. Omeir RL, Arreaza G, Deutsch DG (1999). Identification of two serine residues involved
in catalysis by fatty acid amide hydrolase. Biochem Biophys Res Commun 264(2): 316-320.
111. Omeir RL, Chin S, Hong Y, Ahern DG, Deutsch DG (1995). Arachidonoyl ethanolamide-
[1,2-14C] as a substrate for anandamide amidase. Life sciences 56(23-24): 1999-2005.
112. Osen-Sand A, Catsicas M, Staple JK, Jones KA, Ayala G, Knowles J, et al. (1993). Inhibition
of axonal growth by SNAP-25 antisense oligonucleotides in vitro and in vivo. Nature 364(6436): 445-448.
113. Paglia G, Angel P, Williams JP, Richardson K, Olivos HJ, Thompson JW, et al. (2015). Ion
mobility-derived collision cross section as an additional measure for lipid fingerprinting and identification. Analytical chemistry 87(2): 1137-1144.
114. Park WJ, Kothapalli KS, Lawrence P, Tyburczy C, Brenna JT (2009). An alternate pathway
to long-chain polyunsaturates: the FADS2 gene product Delta8-desaturates 20:2n-6 and 20:3n-3. Journal of lipid research 50(6): 1195-1202.
115. Patricelli M CB (1999). Fatty Acid Amide Hydrolase Competitively Degrades Bioactive
Amides and Esters through a Nonconventional Catalytic Mechanism†. Biochemistry 38(14125-14130).
116. Patricelli MP, Lovato MA, Cravatt BF (1999). Chemical and mutagenic investigations of
fatty acid amide hydrolase: evidence for a family of serine hydrolases with distinct catalytic properties. Biochemistry 38(31): 9804-9812.
117. Pawlosky RJ, Hibbeln JR, Novotny JA, Salem N, Jr. (2001). Physiological compartmental
analysis of alpha-linolenic acid metabolism in adult humans. Journal of lipid research 42(8): 1257-1265.
145
118. Pereira SL, Leonard AE, Mukerji P (2003). Recent advances in the study of fatty acid
desaturases from animals and lower eukaryotes. Prostaglandins, leukotrienes, and essential fatty acids 68(2): 97-106.
119. Perlmann T, Wallen-Mackenzie A (2004). Nurr1, an orphan nuclear receptor with
essential functions in developing dopamine cells. cell and tissue research 318(1): 45-52.
120. Pfenninger KH, Johnson MP (1983). Membrane biogenesis in the sprouting neuron. I.
Selective transfer of newly synthesized phospholipid into the growing neurite. J cell Biol 97(4): 1038-1042.
121. Picq M, Chen P, Perez M, Michaud M, Vericel E, Guichardant M, et al. (2010). DHA
metabolism: targeting the brain and lipoxygenation. Molecular neurobiology 42(1): 48-51.
122. Piomelli D (1994). Eicosanoids in synaptic transmission. Critical reviews in neurobiology
8(1-2): 65-83.
123. Piomelli D, Astarita G, Rapaka R (2007). A neuroscientist's guide to lipidomics. Nature
reviews. Neuroscience 8(10): 743-754.
124. Pitler TA, Alger BE (1994). Depolarization-induced suppression of GABAergic inhibition in
rat hippocampal pyramidal cells: G protein involvement in a presynaptic mechanism. Neuron 13(6): 1447-1455.
125. Posse de Chaves E, Vance DE, Campenot RB, Vance JE (1995). Axonal synthesis of
phosphatidylcholine is required for normal axonal growth in rat sympathetic neurons. J cell Biol 128(5): 913-918.
126. Purvis JM, Clandinin MT, Hacker RR (1982). Fatty acid accretion during perinatal brain
growth in the pig. A model for fatty acid accretion in human brain. Comparative biochemistry and physiology. B, Comparative biochemistry 72(2): 195-199.
127. Qi K, Hall M, Deckelbaum RJ (2002). Long-chain polyunsaturated fatty acid accretion in
brain. Curr Opin Clin Nutr Metab Care 5(2): 133-138.
146
128. Rapoport SI (2003). In vivo approaches to quantifying and imaging brain arachidonic and docosahexaenoic acid metabolism. J Pediatr 143(4 Suppl): S26-34.
129. Rapoport SI (2005). In vivo approaches and rationale for quantifying kinetics and
imaging brain lipid metabolic pathways. Prostaglandins & other lipid mediators 77(1-4): 185-196.
130. Rapoport SI, Ramadan E, Basselin M (2011). Docosahexaenoic acid (DHA) incorporation
into the brain from plasma, as an in vivo biomarker of brain DHA metabolism and neurotransmission. Prostaglandins & other lipid mediators 96(1-4): 109-113.
131. Rapoport SI, Rao JS, Igarashi M (2007). Brain metabolism of nutritionally essential
polyunsaturated fatty acids depends on both the diet and the liver. Prostaglandins, leukotrienes, and essential fatty acids 77(5-6): 251-261.
132. Rashid MA, Katakura M, Kharebava G, Kevala K, Kim HY (2013). N-
Docosahexaenoylethanolamine is a potent neurogenic factor for neural stem cell differentiation. J Neurochem 125(6): 869-884.
133. Read DE, Gorman AM (2009). Involvement of Akt in neurite outgrowth. be llular and
molecular life sciences : CMLS 66(18): 2975-2984.
134. Reddy TS, Sprecher H, Bazan NG (1984). Long-chain acyl-coenzyme A synthetase from
rat brain microsomes. Kinetic studies using [1-14C]docosahexaenoic acid substrate. European journal of biochemistry / FEBS 145(1): 21-29.
135. Rodriguez de Fonseca F, Del Arco I, Bermudez-Silva FJ, Bilbao A, Cippitelli A, Navarro M
(2005). The endocannabinoid system: physiology and pharmacology. Alcohol and alcoholism 40(1): 2-14.
136. Rodriguez OC, Schaefer AW, Mandato CA, Forscher P, Bement WM, Waterman-Storer
CM (2003). Conserved microtubule-actin interactions in cell movement and morphogenesis. Nature cell biology 5(7): 599-609.
137. Rosa AO, Rapoport SI (2009). Intracellular- and extracellular-derived Ca(2+) influence
phospholipase A(2)-mediated fatty acid release from brain phospholipids. Biochimica et biophysica acta 1791(8): 697-705.
147
138. Sanson B, Wang T, Sun J, Wang L, Kaczocha M, Ojima I, et al. (2014). Crystallographic
study of FABP5 as an intracellular endocannabinoid transporter. Acta crystallographica. Section D, Biological crystallography 70(Pt 2): 290-298.
139. Schlicker E, Kathmann M (2001). Modulation of transmitter release via presynaptic
cannabinoid receptors. Trends in pharmacological sciences 22(11): 565-572.
140. Schmid H SP, Natarajan V (1996). The N-acylation-phosphodiesterase pathway and cell
signalling. Chemistry and Physics of Lipids 80: 133-142.
141. Schmid HH (2000). Pathways and mechanisms of N-acylethanolamine biosynthesis: can
anandamide be generated selectively? Chem Phys Lipids 108(1-2): 71-87.
142. Schmid HH, Schmid PC, Natarajan V (1996). The N-acylation-phosphodiesterase pathway
and cell signalling. Chemistry and physics of lipids 80(1-2): 133-142.
143. Schmid PC, Zuzarte-Augustin ML, Schmid HH (1985). Properties of rat liver N-
acylethanolamine amidohydrolase. The Journal of biological chemistry 260(26): 14145-14149.
144. Sheskin T, Hanus L, Slager J, Vogel Z, Mechoulam R (1997). Structural requirements for
binding of anandamide-type compounds to the brain cannabinoid receptor. Journal of medicinal chemistry 40(5): 659-667.
145. Siguira T KS, Sukagawa A, Waku K. (1996). Transacylase-mediated and
phosphodiesterase-mediated synthesis of N-arachidonoylethanolamine, an endogenous cannabinoid-receptor ligand, in rat brain microsomes. Comparison with synthesis from free arachidonic acid and ethanolamine. Eur. J. Biochem. 240: 53-62
146. Snider NT, Nast JA, Tesmer LA, Hollenberg PF (2009). A cytochrome P450-derived
epoxygenated metabolite of anandamide is a potent cannabinoid receptor 2-selective agonist. Molecular pharmacology 75(4): 965-972.
147. Sprecher H (2000). Metabolism of highly unsaturated n-3 and n-6 fatty acids. Biochimica
et biophysica acta 1486(2-3): 219-231.
148
148. Strokin M, Sergeeva M, Reiser G (2003). Docosahexaenoic acid and arachidonic acid release in rat brain astrobytes is mediated by two separate isoforms of phospholipase A2 and is differently regulated by byclic AMP and Ca2+. British journal of pharmacology 139(5): 1014-1022.
149. Sundler R, Akesson B, Nilsson A (1974). Effect of different fatty acids on glycerolipid
synthesis in isolated rat hepatobytes. The Journal of biological chemistry 249(16): 5102-5107.
150. Tai EK, Wang XB, Chen ZY (2013). An update on adding docosahexaenoic acid (DHA) and
arachidonic acid (AA) to baby formula. Food & function 4(12): 1767-1775.
151. Tang BL (2001). Protein trafficking mechanisms associated with neurite outgrowth and
polarized sorting in neurons. J Neurochem 79(5): 923-930.
152. Thors L, Fowler CJ (2006). Is there a temperature-dependent uptake of anandamide into
cells? British journal of pharmacology 149(1): 73-81.
153. Tiger G, Stenstrom A, Fowler CJ (2000). Pharmacological properties of rat brain fatty acid
amidohydrolase in different subcellular fractions using palmitoylethanolamide as substrate. Biochem Pharmacol 59(6): 647-653.
154. Tocher DR, Leaver MJ, Hodgson PA (1998). Recent advances in the biochemistry and
molecular biology of fatty acyl desaturases. Progress in lipid research 37(2-3): 73-117.
155. Ueda N KY, Yamamoto S. (1995). Partial Purification and Characterization of the Porcine
Brain Enzyme Hydrolyzing and Synthesizing Anandamide. The Journal of biological chemistry 270(40): 23823–23827.
156. Ueda N, Puffenbarger RA, Yamamoto S, Deutsch DG (2000). The fatty acid amide
hydrolase (FAAH). Chem Phys Lipids 108(1-2): 107-121.
157. Ueda N, Tsuboi K, Uyama T (2010). Enzymological studies on the biosynthesis of N-
acylethanolamines. Biochimica et biophysica acta 1801(12): 1274-1285.
149
158. Uyama T, Ikematsu N, Inoue M, Shinohara N, Jin XH, Tsuboi K, et al. (2012). Generation of N-acylphosphatidylethanolamine by members of the phospholipase A/acyltransferase (PLA/AT) family. The Journal of biological chemistry 287(38): 31905-31919.
159. van Meer G, Lisman Q (2002). Sphingolipid transport: rafts and translocators. The
Journal of biological chemistry 277(29): 25855-25858.
160. van Meer G, Voelker DR, Feigenson GW (2008). Membrane lipids: where they are and
how they behave. Nature reviews. Molecular cell biology 9(2): 112-124.
161. Vance JE, Vance DE (2004). Phospholipid biosynthesis in mammalian cells. Biochemistry
and cell biology = Biochimie et biologie cellulaire 82(1): 113-128.
162. Walker JM, Huang SM, Strangman NM, Tsou K, Sanudo-Pena MC (1999). Pain
modulation by release of the endogenous cannabinoid anandamide. Proceedings of the National Academy of Sciences of the United States of America 96(21): 12198-12203.
163. Wallen-Mackenzie A, Mata de Urquiza A, Petersson S, Rodriguez FJ, Friling S, Wagner J,
et al. (2003). Nurr1-RXR heterodimers mediate RXR ligand-induced signaling in neuronal cells. Genes & development 17(24): 3036-3047.
164. Wang J, Ueda N (2009). Biology of endocannabinoid synthesis system. Prostaglandins &
other lipid mediators 89(3-4): 112-119.
165. Welser-Alves J (2015). The Advantages and Difficulties of Working with Primary be lls. In:
The American Society for cell BiologyThe American Society for cell Biology.
166. Wenk MR (2005). The emerging field of lipidomics. Nature reviews. Drug discovery 4(7):
594-610.
167. Williams CM, Kirkham TC (1999). Anandamide induces overeating: mediation by central
cannabinoid (CB1) receptors. Psychopharmacology 143(3): 315-317.
168. Williard DE, Harmon SD, Kaduce TL, Preuss M, Moore SA, Robbins ME, et al. (2001).
Docosahexaenoic acid synthesis from n-3 polyunsaturated fatty acids in differentiated rat brain astrobytes. Journal of lipid research 42(9): 1368-1376.
150
169. Wilson D B PSM, Majerus P W. (1982). Discovery of an Arachidonoyl Coenzyme A
Synthetase in Human Platelets. THE JOURNAL OF BIOLOGICAL CHEMISTRY 257(7): 3510-3515.
170. Wilson RI, Kunos G, Nicoll RA (2001a). Presynaptic specificity of endocannabinoid
signaling in the hippocampus. Neuron 31(3): 453-462.
171. Wilson RI, Nicoll RA (2001b). Endogenous cannabinoids mediate retrograde signalling at
hippocampal synapses. Nature 410(6828): 588-592.
172. Yan S, Yang XF, Liu HL, Fu N, Ouyang Y, Qing K (2015). Long-chain acyl-CoA synthetase in
fatty acid metabolism involved in liver and other diseases: an update. World journal of gastroenterology 21(12): 3492-3498.
173. Yang HC, Mosior M, Johnson CA, Chen Y, Dennis EA (1999). Group-specific assays that
distinguish between the four major types of mammalian phospholipase A2. Analytical biochemistry 269(2): 278-288.
174. Yang R, Fredman G, Krishnamoorthy S, Agrawal N, Irimia D, Piomelli D, et al. (2011).
Decoding functional metabolomics with docosahexaenoyl ethanolamide (DHEA) identifies novel bioactive signals. The Journal of biological chemistry 286(36): 31532-31541.
175. Yang S, Edman LC, Sanchez-Alcaniz JA, Fritz N, Bonilla S, Hecht J, et al. (2013).
Cxcl12/Cxcr4 signaling controls the migration and process orientation of A9-A10 dopaminergic neurons. Development 140(22): 4554-4564.
176. Youdim (2000). Essential Fatty acids and the brain: possible health implications. Int.
journal of Devel. Neurosciences.
177. Youdim KA, Martin A, Joseph JA (2000). Essential fatty acids and the brain: possible
health implications. International journal of developmental neuroscience : the official journal of the International Society for Developmental Neuroscience 18(4-5): 383-399.
![Amazon Web Services · 2020. 9. 14. · $58 $38 $48 $68 $65 Tel: 3704 7177 Tel: 3709 8455 Tel: 2310 2818 $35 $42 C) $48 $38 $45 $50 $48 $65 C] $38 C] $48 C] $68 C] $48 C] $58 C] $38](https://img.pdfslide.us/doc/110x75/60f8ecca7613ef7a9e5913dc/amazon-web-services-2020-9-14-58-38-48-68-65-tel-3704-7177-tel-3709.jpg)
