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N-acylethanolamines as novel alcohol dehydrogenase 3 substrates

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Title:
N-acylethanolamines as novel alcohol dehydrogenase 3 substrates
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English
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Ivkovic, Milena
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University of South Florida
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Subjects / Keywords:
Enzyme
Kinetics
N-acylglycinals
N-acylglycines
Amides
Dissertations, Academic -- Chemistry -- Masters -- USF   ( lcsh )
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non-fiction   ( marcgt )

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Abstract:
ABSTRACT: N-Acylethanolamines (NAEs) are a class of fatty acid amides that act as important mammalian signaling molecules. N-Arachidonoylethanolamine is the best-studied representative and is one of the endogenous ligands for endocannabinoid receptors. NAEs play a role in the regulation of appetite, act as anti-inflammatory and analgesic agents, and are thought to have a neuroprotective function as well. They have been proposed to also serve as precursors to N-acylglycines (NAGs). N-Acylglycinals are likely to be intermediates between the NAEs and the NAGs. The sequential actions of a putative fatty alcohol dehydrogenase and a putative fatty aldehyde dehydrogenase are thought to affect the NAD+-dependent oxidation of the NAEs to the NAGs. NAGs, in turn, serve as precursor in the biosynthesis of primary fatty acid amides (PFAMs), another class of mammalian regulatory molecules. Alcohol dehydrogenase 3 (ADH3), an enzyme known to oxidize mid- and long-chain alcohols to aldehydes, was evaluated for its potential in oxidation of NAEs to N-acylglycinals. In order to evaluate the possibility of ADH3 involvement in NAE metabolism, variable chain length NAEs were synthesized and evaluated as substrates for bovine liver ADH3. NAEs were oxidized by ADH3 in the presence of NAD+, yielding the corresponding N-acylglycinals. Vsubfield Max/Ksubfield M values for assayed NAEs were low relative to cinnamyl alcohol, one of the preferred substrates for ADH3. Our data suggest that the ADH-mediated oxidation of NAEs could occur in vivo, but that ADH3 is unlikely to be the in vivo catalyst.
Thesis:
Thesis (M.S.)--University of South Florida, 2008.
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Includes bibliographical references.
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by Milena Ivkovic.
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N -Acylethanolamines as Novel Alcohol Dehydrogenase 3 Substrates by Milena Ivkovic A thesis submitted in partial fulfillment of the requirements for the degree of Master of Science Department of Chemistry College of Arts and Sciences University of South Florida Major Professor: David J. Merkler, Ph.D. Wayne C. Guida, Ph.D. Kirpal S. Bisht, Ph.D. Date of Approval: November 18, 2008 Keywords: enzyme, kinetics, N -acylglycinals, N -acylglycines, amides Copyright 2008, Milena Ivkovic

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I would like to dedicate this work to my parents, Ljubica and Milo rad, and my brother, Goran. Without their love and support I would not be where I am today.

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Acknowledgements I would like to thank Dr. Da vid Merkler for giving me the opportunity to work in his lab, and for helping to guide my project to a successful completion. I would also like to thank my commi ttee members, Dr. Wayne Guida and Dr. Kirpal Bisht, for their help ful suggestions and advice. I would like to acknowledge E. Will Lowe who performed the modeling experiments, and helped me with advice and suggestions throughout the project. My thanks to Jacob Shafer for synthesis of N -octanoylglycine, and for his constant friendship, advice and support. Thanks to Emma Farrell for her assistance with GC-MS, and her kindness and friendship. Thanks to Neil McIntyre for first inviting me to work in the Merkler lab. I would also like to acknowledge othe r Merkler group members, Sumit Handa, Zhenming An, and Shikha Mahajan for thei r help and support throughout my graduate school experience.

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i Table of Contents List of Tables ....................................................................................................................iii List of Figures....................................................................................................................iv List of Schemes................................................................................................................ ..vi Abstract....................................................................................................................... ......vii Chapter 1 Introduction....................................................................................................... 1 N -Acylethanolamines (NAEs)..................................................................................1 Alcohol Dehydrogenases (ADHs)............................................................................4 Alcohol Dehydrogenase 3 (ADH3).............................................................6 Chapter 2 Material and Methods.......................................................................................9 Enzyme Purification................................................................................................. .9 A. Ammonium Sulfate Precipitation............................................................9 B. DEAE-Cellulose Chromatography........................................................10 C. Mimetic Blue II Chromatography.........................................................10 D. Blue Dextran Agarose Chromatography...............................................10 E. Enzyme Activity Assay.........................................................................11 Syntheses.................................................................................................................11 A. Synthesis of NAEs................................................................................11 B. Synthesis of [1,2-13C]N -Octanoylethanolamine................................12 C. Synthesis of N -Acylglycinals................................................................13 D. Synthesis of N -Octanoylglycine............................................................14 E. Preparation of N -Octanoylglycinal semicarbazone...............................15 Kinetic Assays of NAE Substrate Activity with ADH3.........................................15 A. Spectrophotometric NADH Assay........................................................15 B. MTS-Formazan NADH Assay..............................................................15 Modeling Experiments............................................................................................16

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ii Product Characterization Experiments...................................................................17 A. HPLC Separation Experiments.............................................................17 B. GC-MS Experiments.............................................................................17 C. Trapping N -Acylglycinals with Semicarbazide....................................19 D. N -Acylglycinal Semicarbazone Characterization by 13C NMR............19 Chapter 3 Results and Discussion...................................................................................21 ADH3 Purification..................................................................................................21 Kinetics of ADH3-catalyzed NAE Oxidation.........................................................22 Docking Experiments..............................................................................................24 Product Characterization.........................................................................................26 Conclusion..............................................................................................................32 References Cited............................................................................................................... .34 Appendices.........................................................................................................................39 Appendix A: NMR Spectra.....................................................................................40 Appendix B: HPLC Spectra....................................................................................54 Appendix C: GC-MS Spectra.................................................................................56

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iii List of Tables Table 1 Kinetic Constants for Shortand Medium-chain NAEs....................................23 Table 2 Relative Rates for Mediumand Long-chain NAEs..........................................23 Table 3 Kinetic Constants for Aromatic NAEs and Cinnamyl Alcohol.........................24 Table 4 Relative Binding Energies and KM Values for Selected NAEs and Cinnamyl Alcohol..............................................................................................25

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iv List of Figures Figure 1 NAE Biosynthesis Reaction...............................................................................2 Figure 2 NAE Degradation Pathway................................................................................2 Figure 3 Proposed Pathway for Bi osynthesis of PFAMs from NAEs..............................3 Figure 4 Aldehyde Dismutation Reaction.........................................................................5 Figure 5 Formaldehyde Detoxification Pathway..............................................................6 Figure 6 ADH3 Dimer......................................................................................................7 Figure 7 MTS-Formazan Assay for NADH Detection...................................................16 Figure 8 SDS-PAGE of Purified Bovine Liver ADH3...................................................21 Figure 9 A Representiative Michaelis-Ment en Plot for One of the Assayed NAEs.......22 Figure 10 N -Hexanoylethanolamine Docked into the Active Site of Human ADH3.....25 Figure 11 HPLC Separation of ADH3-catalyzed N -Benzoylethanolamine Oxidation.........................................................................................................26 Figure 12 HPLC Separation of ADH3-catalyzed Cinnamyl Alcohol Oxidation............27 Figure 13 GC-MS Spectrum of Cinnamyl Aldehyde O -PFB-oximes............................28 Figure 14 GC-MS Spectrum of N,O-di-TMSN -Benzoylglycine...................................29 Figure 15 Semicarbazone Formation Reaction...............................................................30 Figure 16 HPLC Separation of ADH3-catalyzed N -Benzoylethanolamine Oxidation in the Presence of Semicarbazide..................................................30 Figure 17 13C NMR Spectrum of ADH3-catalyzed [1,2-13C]N -Octanoylethanolamine Oxidation in the Presence of Semicarbazide.................................................31 Figure 18 13C NMR Spectrum of ADH3-catalyzed [1,2-13C]N -Octanoylethanolamine Oxidation without Semicarbazide..................................................................32 Figure A-1 1H and 13C Spectra of N -Butyrylethanolamine in DMSO-d6.......................40 Figure A-2 1H and 13C Spectra of N -Hexanoylethanolamine in DMSO-d6....................41

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v Figure A-3 1H and 13C Spectra of N -Octanoylethanolamine in DMSO-d6.....................42 Figure A-4 1H and 13C Spectra of N -Decanoylethanolamine in DMSO-d6....................43 Figure A-5 1H and 13C Spectra of N -Lauroylethanolamine in DMSO-d6.......................44 Figure A-6 1H and 13C Spectra of N -Myristoylethanolamine in DMSO-d6....................45 Figure A-7 1H and 13C Spectra of N -Oleoylethanolamine in DMSO-d6........................46 Figure A-8 1H and 13C Spectra of N -Benzoylethanolamine in DMSO-d6......................47 Figure A-9 1H and 13C Spectra of [1,2-13C]N -Octanoylethanolamine in DMSO-d6..48 Figure A-10 1H and 13C Spectra of N -Benzoylglycinal in CDCl3..................................49 Figure A-11 1H and 13C Spectra of N -Octanoylglycinal in CDCl3.................................50 Figure A-12 1H and 13C Spectra of N -Octanoylethanolamine semicarbazone in DMSO-d6...................................................................................................51 Figure A-13 1H and 13C Spectrum of N -Octanoylglycine in DMSO-d6.........................52 Figure A-14 13C Spectrum of [1,2-13C]N -Octanoylethanolamine Control Sample without ADH3............................................................................................53 Figure B-1 HPLC Separation of N -Benzoylethanolamine and its Derivatives...............54 Figure B-2 HPLC Separation of Cinna myl Alcohol and it s Derivatives........................55 Figure B-3 HPLC Separation of Semicarbazide and N -Octanoylglycinal semicarbazone...............................................................................................55 Figure C-1 GC-MS Spectra of Cinnamyl Aldehyde O -PFB-oxime Standard................56 Figure C-2 GC-MS Spectra of TMS-derivatized N -Benzoylglycine Standard..............57 Figure C-3 GC-MS Spectru m of N-Benzoylglycinal O -PFB-oxime Standard..............58

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List of Schem es Scheme 1. Synthesis of NAEs...........................................................................................12 Scheme 2. Synthesis of [1,2-13C]N -Octanoylethanolamine...........................................12 Scheme 3. Synthesis of N -Acylglycinals...........................................................................14 Scheme 4. PFBHA Derivatization Reaction......................................................................18 Scheme 5. BSTFA Derivatization Reaction......................................................................18 vi

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vii N -Acylethanolamines as Novel Alcohol Dehydrogenase 3 Substrates Milena Ivkovic ABSTRACT N -Acylethanolamines (NAEs) are a class of fatty acid amides that act as important mammalian signaling molecules. N -Arachidonoylethanolamine is the best-studied representative and is one of the endogenous ligands for e ndocannabinoid receptors. NAEs play a role in the regulation of appetite, ac t as anti-inflammatory and analgesic agents, and are thought to have a neur oprotective function as well. They have been proposed to also serve as precursors to N -acylglycines (NAGs). N -Acylglycinals are likely to be intermediates between the NAEs and the NAGs. The sequential actions of a putative fatty alcohol dehydrogenase and a putative fatty aldehyde dehydroge nase are thought to affect the NAD+-dependent oxidation of the NAEs to the NAGs. NAGs, in turn, serve as precursor in the biosynthesis of primary fa tty acid amides (PFAMs), another class of mammalian regulatory molecules. Alcohol dehydrogenase 3 (ADH3), an enzyme known to oxidize midand long-chain alcohols to aldehydes, was ev aluated for its potential in oxidation of NAEs to N -acylglycinals. In order to evaluate the possibility of ADH3 involvement in NAE metabolism, variable chain length NAEs were synthesized and evaluated as substrates for bovine liver ADH3. NAEs were oxidized by ADH3 in the presence of NAD+, yielding the corresponding N -acylglycinals. VMax/KM values for assayed NAEs were low relative to cinnamyl alcohol, one of the preferred substr ates for ADH3. Our data suggest that the ADH-mediated oxidation of NAEs could occur in vivo but that ADH3 is unlikely to be the in vivo catalyst.

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Chapter I Introduction N-Acylethanolamines (NAEs ) N -Acylethanolamines (NAEs) belong to th e class of fatty acid amides, which also includes primary fatty acid amides (PFAMs), N -acyldopamines (NADAs), N -acylamides (NAMs) and N -acylamino acids (NAAs) (1). They are the most widely studied group of fatty acid amides and represent an important class of mammalian signaling molecules. NAEs have been isolated from the brain as well as a variety of other tissues (1), and have been associated with a number of important processes, such as appetite control, regulation of energy metabolism, and anti-inflammatory response (1-3). N Arachidonoylethanolamine (anandamide) is an endogenous ligand for cannabinoid receptors (CB1 and CB2), and as a result, it is the most extensively studied of all NAEs. Anandamide plays an important role in th e stimulation of appe tite, reduction of body temperature and movement, and also has anal gesic properties (1-4). It is thought that most of these effects of anandamide are mediated by its binding to CB1 receptor. Other long-chain NAEs whose functions have been established also play important roles in the body. N -Palmitoylethanolamine is a well-known anti-inflammatory agent, and also has analgesic properties (1, 2, 5). N -Oleoylethanolamine inhibits food intake through binding to PPAR (peroxisome proliferat or-activated receptor ) and GPR119 (G-protein coupled receptor), there by acting in opposition to anandamide (1, 3). N -Stearoylethanolamine exhibits similar effects as anandamide with regard to analgesia, body temperature and motility, although it is unable to bind to cannabinoid receptors (1). It has also recently been found to possess an ti-inflammatory properties (6). Long-chain NAEs, including N -stearoylethanolamine and N -palmitoylethanolamine, have been thought to simulate the effects of anandami de by acting as entourage compounds (7-8). All long chain NAEs are degraded by fatty acid amide hydrolase (FAAH), and entourage NAEs compete with anandamide for binding to FAAH, thereby effectively 1

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reducing its rate of degrada tion and prolonging its effects in the body. NAEs have also been postulated to have a neuroprotective acti on, as their levels are increased in injured neurons (4). Increase in NAE levels followi ng cell stress and injury has also been found in other organs, such as heart and testes, suggesting they m ay play a similar role in these tissues as well (2). Primary route for NAE synthe sis is by the action of N acylphosphatidylethanolamine-specific phospholipase D (NAPE-PLD), which catalyzes the cleavage of N -acylphosphatidylethanolamine (NAPE) into NAE and phosphatidic acid (PA) (Fig. 1). NAE degradation to fa tty acid and ethanolamine is accomplished primarily through the action of fatty aci d amide hydrolase (FAAH), although another enzyme capable of doing the same chemistry, N -acylethanolamine-hydrolyzing acid amidase (NAAA), has been discovered recently (9 -10) (Fig. 2). Altern atively, it has been proposed that NAEs could act as precursors in the biosynthetic pathway of PFAMs (1, 11), a related class of fatt y acid amides (Fig 3). O O O R1 O O R2 P O O OHN R3 O NAPE-PLD O O O R1 O O R2 P OO ON H R3 O HO + NAPE( N -Acylphoshatidylethanolamine) PA(Phosphatidicacid) NAE( N -Acylethanolamine) Figure 1. NAE biosynthesis reaction. N H R O HO NAE( N -Acylethanolamine) FAAH/NAAA H2OHO R O NH2 HO + Fattyacid Ethanolamine Figure 2. NAE degradation pathway. 2

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N H R O HO NAE( N -Acylethanolamine) FADH(Fattyalcoholdehydrogenase) NAD+NADHN H R O H N -AcylglycinalO FALDH(Fattyaldehydedehydrogenase) NAD+NADHN H R O HO NAG( N -Acylglycine)O PAM (Peptidylglycine-alpha-amidatingmonooxygenase) 2Ascorbate+O22Semihydroascorbate+H2OH2N R O PFAM(Primaryfattyacidamide) glyoxylate Figure 3. Proposed pathway for biosynthesis of PFAMs from NAEs. PFAMs were first identified in human plasma (12), but have since been isolated from brain and cerebrospinal fluid as well. Like NAEs, they function as signaling and regulatory molecules in the body. The best -studied PFAM is oleamide, which is important in regulation of sleep, memory, body temperature and movement (1). Other 3

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naturally occurring PFA Ms include erucamide, linoleamide and elaidamide (1). Initially proposed biosynthetic route to PFAMs was through reaction between fatty acid and ammonia, catalyzed by FAAH (13). Howe ver, although this chemistry occurs in vitro it requires relatively high pH and high ammonia c oncentration, making it unlikely that it is the primary synthetic pathway in vivo There are two other po ssible routes to PFAM biosynthesis. First route involves cytochrome c-catalyzed amidation of fatty acyl-CoA thioester by ammonia (1, 14-15). The other ro ute is by the action of peptidylglycineamidating monooxygenase (PAM) on N -acylglycines (NAG), resu lting in production of PFAMs and glyoxylate (Fig. 3). NAGs are well es tablished as good substrates for PAM, but their biosynthetic pathway is unclear. Th ere is evidence that NAEs could serve as precursors to NAGs through the sequential actions of a putative fatty alcohol dehydrogenase (FADH) and a putative fatty aldehyde dehydrogenase (FALDH) (Fig. 3). Support for this pathway comes from experiments in which 14C-labeled N arachidonoylglycine was detected in liv er cells that were treated with 14C-labeled anandamide (11). These experi ments strongly suggest that th ere is a set of liver ADH and ALDH enzymes capable of catalyzing the conversion of NAEs to NAGs. Alcohol Dehydrogenases (ADHs) Alcohol dehydrogenases (ADHs) comprise a fam ily of NAD+-dependent, Zncontaining, dimeric enzymes re sponsible for oxidation of alcohols to aldehydes. ADH dimers are composed of identical subunits, each containing two Zn ions, one of which plays a structural role, and one which is i nvolved in catalysis. Each subunit is composed of two domains: a coenzyme binding domain, located close to the interface with other subunit and a catalytic domain, located towa rd other ends of the dimer (Fig. 6). ADH enzymes are divided into six classes based on their structure, substrate preference, sensitivity to inhibitors, and electrophoretic mobility. Class I ADH (ADH1) enzymes are expressed in all vertebrates, (16) and in humans they are present in the liver, kidney, lung, mucosa of the stomach and lower digestive tract (17). This class contains a number of isozymes, created through the different combinations of th ree subunits designated as , and Among ADHs, ADH1 4

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enzym es have lowest KM values (< 5mM) and highest kcat values for ethanol, indicating that their primary role is metabolism of etha nol to acetaldehyde. They are also the most strongly inhibited by 4-methylpy razole, and have the highest isoelectric point of all ADH enzymes (17). In addition to ethanol oxidati on, ADH1 isozymes are also able to oxidize retinol, and possess aldehyde dismutase activ ity, converting aldehydes to carboxylic acids and alcohols (Fig. 4) (18). Class II ADH (ADH2) is expressed exclusiv ely in liver, and shares some of the characteristics with ADH1. It has a higher KM and lower kcat value for ethanol than ADH1, but it is likely also involved in etha nol metabolism. It is also inhibited by 4methylpyrazole, but to a smaller extent than ADH1, and it has a lower pI than class I enzymes. Like ADH2, it is capable of utilizing retinol as a substrate, and it has been shown to have aldehyde dismutase activity (18). E-NAD+E-NADH RCHO ROH RCOOH RCHO E+NAD+ E+NADH Figure 4. Aldehyde dismutation reaction. Class IV ADH (ADH4) is found in the muco sa of the stomach and upper digestive tract, and therefore may play a role in the fi rst-pass metabolism of ethanol (19). It shares many features with ADH2 similar KM value for ethanol, similar pI, and similar level of inhibition by 4-methylpyrazole. However, its turnover values for both ethanol and retinol are higher, and it has been suggested that ADH4 is the primary enzyme for oxidation of retinol to retinal (20-22). Class V ADH (ADH5) has been charact erized on cDNA and mRNA level in humans (16, 23), but remains largely unstudied. Class VI (ADH6) has been isolated as cDNA from deer mouse and rat (16, 20, 24), but has also not b een investigated to greater effect. The functions of these cla sses therefore remain largely unknown. 5

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Alcohol Dehydrogenase 3 (ADH3) Class III alcohol dehydrogenase (ADH3) is found in virtually all eukaryotes and prokaryotes and represents the ancestral enzyme fro m which all the other ADHs originated. It was first is olated from human liver in 1981 (25), and it was fully characterized in 1984 (26). S hortly thereafter, it was recogni zed that ADH3 is identical to the previously characteriz ed glutathione-dependent formaldehyde dehydrogenase (2728), which catalyzes the oxidation of S-(hydroxymethyl)-glutathione (HMGSH) (formed through non-enzymatic conjugation of formal dehyde to glutathione (GSH)) (Fig. 5). Together with S-formylglutathione hydrolase, ADH3 forms a major formaldehyde detoxification pathway in the body (Fig 5). Figure 5. Formaldehyde detoxification pathway. Ethanol is a very poor substrate for ADH3, and saturation is never observed, precluding the determination of KM and VMax values. Unlike all th e other characterized ADHs, ADH3 is not inhibited by 4-methylpyrazole, and cannot oxidize retinol. It also lacks the aldehyde dismutase activity of ADH1 and ADH2 (18). Besides HMGSH, the best substrates for ADH3 are mediumand l ong-chain alcohols (5 carbons and up), and -hydroxy fatty acids, although it also able to oxidize 20-hydroxyleukotriene B4 (29). This preference for larger substrates can be explained by the differences in the active site size and structur e between ADH3 and other ADH enzymes characterized to date. In all ADH enzymes, active site is positioned in the cleft between the coenzyme binding domain and the catalytic domain of the subunit (Fig. 6). In ADH1 and ADH4, 6

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binding of the coenzyme induces a confor matio nal change whereby cat alytic unit rotates by ~10o toward the coenzyme-binding domain, th ereby converting the enzyme from the open to the closed confor mation (30-35). Closure of the active site enables tighter binding for the smaller substrates, accounting for the preference these enzymes have for shorter alcohols. Because NAD+ binding is required in order to achieve effective substrate binding, these enzymes display an ordered bi-bi mechanism (31-32, 36). Figure 6. ADH3 dimer (adapted from ref. 30). Subunits are colored blue and green. Coenzyme binding domains are colored light blue and light green, and catalytic domains are dark blue and dark green. Catalytic Zn ions are colored red, and structural Zn ions are shown in grey. Bound NAD+ molecules are shown in purple. Yellow areas represen t important secondary structures in the proximity of the active sites. In contrast, the ADH3 active site displays a semi-open configuration that is inbetween the fully open and fully close d conformations exhibited by ADH1 (30-32). In addition to this, coenzyme binding resu lts in only small conformational changes between the two domains, leaving a large, solvent-exposed active site approximately twice the size of the ADH1 active site (30) Substrate binding is th erefore not dependent on prior NAD+ binding, and, as a result, ADH3 has a random bi-bi mechanism (36). Having a large active site means that ADH3 is capable of accommodating bulky substrates, like long chain alcohols, -hydroxy fatty acids and HMGSH. It also accounts for its poor activity toward th e smaller substrates, such as ethanol, which are unable to form the bonding interactions they make in other ADHs. In addition to the open active 7

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site, sm aller substrates lack the stabili zation provided by the loop formed by residues 112 to 120 in ADH1, which is replaced by an -helix in ADH3 which fails to provide any binding support (30). The well-established preference of ADH3 for large, hydrophobic alcohol substrates suggests that it could act as the putative FADH catalyzing the oxidation of NAEs to N -acylglycinals. It is also the most ubi quitous of all the ADH enzymes, and it is the sole ADH representative so far characteri zed in the brain (37), where abundance of NAEs is found. Its substrate preference combin ed with its presence in the tissues where NAEs have been found, led us to investigat e ADH3 as a potential FADH, acting in the first step of the proposed PFAM biosyntheti c pathway to catalyze the oxidation of NAEs to N -acylglycinals (Fig. 3). 8

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Chapter 2 Materials and Methods Enzyme Purification Bovine liver ADH3 was purified foll owing the modified procedure of Pourm otabbed et al. (38). Purification was accomplished by performing ammonium sulfate precipitation, followed by DEAE-cellulose, mimetic blue II, and blue dextran agarose chromatography. A. Ammonium Sulfate Precipitation Fresh bovine liver was obtained from Central Packing Co Inc, (Center Hill, FL), cut into parts, and frozen at 80 oC. Bovine liver portion weighing 265 g was thawed out, chopped into small pieces, combined with 795 ml of 50 mM potassium phosphate pH 7.5, 5 mM -mercaptoethanol (BME), and a cocktail of protase inhibitors (1g/ml leupeptin, 1 g/ml pepstatin A, and 1 g/ml aprotinin), and blended for 3 x 30 seconds in a Waring blender at high setting. Blended mixture was centrifuged in a Sorva ll RC 6 centrifuge at 20,000 x g and 4 oC for 40 minutes. Pellet was then discarded and supernatant centrifuged at 32,000 x g and 4 oC for 30 minutes. Reddish-brown supernatant (670 ml) was obtained following the second centrif ugation, and taken to 45% saturation by addition of 173 g of ammonium sulfate. Finely ground ammonium sulfate was added slowly over a 1-hour period with continuous stirring, and pH of the solution was maintained at 7.4 by addition of concentrated NH4OH. Following salt addition, solution was left to equilibrate for 2 hours without stirring. Equilibrated solution was centrifuged at 47,800 x g for 30 minutes. Recovered supern atant (600 ml) was then taken to 67% saturation by addition of 84 g of ammonium sulfate. As previously, ammonium sulfate was added slowly over 1 hour and pH was maintained by addition of concentrated NH4OH. Following equilibration, solution was centrifuged at 47,800 x g for 30 minutes. Supernatant was discarded and pellet dissolv ed in minimum amount of 10 mM Tris pH 9

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7.6 and 5 mM BME solution. Sam ple was then dialyzed in the same buffer in order to remove the salt. B. DEAE-Cellulose Chromatography Following dialysis, sample was loaded onto a DEAE cellulose column (2 x 40 cm) previously equilibrated with 10 mM Tris pH 7.6 and 5 mM BME. Column was washed with running buffer until A280 was reduced to 0.2, and el uent was collected in 15 ml fractions. ADH3 was eluted off the column by running a li near pH gradient with 500 ml of running buffer (10 mM Tris pH 7.6) a nd 500 ml of 10 mM Tr is pH 7.0. Fractions were assayed for protein content by meas uring absorbance at 280 nm, while enzyme activity was assayed by following NADH producti on using cinnamyl alcohol as substrate. Fractions with highest specific activity were pooled t ogether and dialyzed into solution of 10 mM Tris pH 8.0 and 2 mM dithiothreitol (DTT). C. Mimetic Blue II Chromatography Dialyzed sample was loaded onto a Mime tic Blue II affinity column (1.5 x 20 cm) that was previously equilibrated with 10 mM Tris pH 8.0 and 2 mM DTT. Column was washed with the same buffer and flow rate was adjusted to 1 ml/min. Eluent was collected in 4 ml fractions. Protein content was monitored by measuring absorbance of fractions at 280 nm. Once A280 was below 0.2, a linear gradient of 0-20 mM NAD+ (total volume 200 ml) was applied to the column in order to elute ADH3. Fractions were assayed for enzyme activity by following NADH production at 340 nm, with cinnamyl alcohol as substrate. Fractions with highest spec ific activity were pooled together and dialyzed into 0.5 mM Tris pH 7.0 and 5 mM BME. D. Blue Dextran Agarose Chromatography Following dialysis to remove NAD+, sample was loaded onto a Blue Dextran agarose column (5 ml), that had been equi librated with 0.5 mM Tris pH 7.0 and 5 mM BME. Column was washed with the same buffer and the eluent was collected in 3 ml fractions, which were assayed for protein co ntent and enzymatic activity as described before. Fractions with the hi ghest specific activity were an alyzed for purity using SDS10

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PAGE, and t hose containing only a single protein band corresponding to the weight of an ADH3 subunit (~40 kDa) were pooled together Protein concentration of purified ADH3 was determined using the Bradford assay, a nd concentrated enzyme was stored at -80 oC. E. Enzyme Activity Assay Enzyme activity was evaluated by following NADH production at 340 nm, with cinnamyl alcohol as the substrate. Assays were done in 100 mM glycine pH 10, 5 mM cinnamyl alcohol, and 2.5 mM NAD+ at 37 oC. NADH production was followed at 340 nm using JASCO V-530 UV-Vis spectrophoto meter. When scanning crude enzyme fractions, which are know to include ADH1 and ADH2, assay also included 3 mM pyrazole, a know inhibitor of these two enzymes. In addition to this, rates for controls without the substrate were obt ained and subtracted from th e reaction rates in order to eliminate false positives. Syntheses A. Synthesis of NAEs A number of N -acylethanolamines were available commercially in high purity and these were obtained and used without further purification. N -Arachidonoylethanolamine and N -linoleoylethanolamine were purchas ed from Cayman Chemical Company, N acetylethanloamine was obtaine d form Sigma-Aldrich, and N -propionylethanolamine was purchased from TCI. The remaining N -acylethanolamines were synthesi zed as described in Jonsson et al. (7-8) (Scheme 1). Briefly, an excess of et hanolamine (~ tenfold) was added to 15-20 ml of anhydrous CH2Cl2 and placed in an ice-bath. Ac yl chloride (0.012 moles) was added drop-wise to the solution while stirring and under N2. Upon complete addition of the chloride, reaction mixture was removed from ice and left stirring at room temperature for another 12 hours. Midand long-chain NAEs (octanoyl and l onger) were purified by recrystallization in ethanol/w ater. Short-chain NAEs (but yryl, hexanoyl and benzoyl) were purified using a silica column run in ethy l acetate: methanol (80:20). The yields for the synthesized NAEs were as follows: N -butyrylethanolamine 48.3 %, N 11

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hexanoylethanolam ine 20.6 %, N -benzoylethanolamine 28.2 %, Noctanoylethanolamine 44.5 %, N -decanoylethanolamine 77.4 %, Nlauroylethanolamine 77.6 %, N -myristoylethanolamine 73.4 %, N -oleoylethanolamine 84.5 %. Purity and identity of synthesized NAEs were confirmed by 1H and 13C NMR on iNOVA 400 MHz (see Figures A1-A8). All ac yl chlorides and ethanolamine were obtained from Sigma-Aldrich, and used without further purification. Anhydrous CH2Cl2 was purchased from EMD. R Cl O H2N OH + R N H O OH N2,CH2Cl2(anhyd.) 12hours Scheme 1. Synthesis of NAEs. B. Synthesis of [1,2-13C]N -Octanoylethanolamine Uniformly 13C-labeled ethanolamine HCl was dissolved in CH3CN and a 10-fold molar excess of triethylamine (TEA) under N2. One equivalent of octanoyl chloride was added drop-wise, and the reaction mixture was left to stir at room temperature for 12 hours (Scheme 2). Reaction progress was mon itored by TLC (ethyl acetate: methanol (80:20 (v/v))). Once reaction was > 90% co mplete, solvent was removed, and residue was redissolved in the same mob ile phase used for TLC. [1,2-13C] N Octanoylethanolamine was purified by silica co lumn chromatography in 53 % yield, and characterized by 1H and 13C NMR spectra obtained on the Va rian iNOVA 400 (Fig. A-9). Scheme 2. Synthesis of [1,2-13C]N -Octanoylethanolamine. 13C labeled atoms are marked by *. 12

PAGE 23

C. Synthesis of N -Acylglycinals N -Benzoylglycinal and N -octanoy lglycinal were prepared using the modified procedure of Brown (39) (Scheme 3). Am inoacetaldehyde diethyl acetal (50 mmoles) was dissolved in 75 ml of diethyl ether and pl aced in an ice-bath. Next, one equivalent (5 ml of 15 M) KOH was added to the solution. On e equivalent of acyl chloride and another two equivalents of KOH were simultaneously added drop-wise to the reaction mixture. Once addition was complete, reaction was le ft stirring on ice for another 3 hours. Reaction progress was monitored by TLC (h exanes: ethyl acetate (1:1)). Ether and aqueous layers were separate d, and ether layer was washed with 5 x 30 ml of saturated NaCl, and dried with MgSO4. Solvent was removed under vacuum to obtain N acylglycinal diethyl acetal. Product pur ity and identity were confirmed by 1H and 13C NMR on Burker DPX 250. N -Benzoylglycinal and N -octanoylglycinal were obtained by acid-catalyzed deprotection of the corresponding N -acylglycinal diethyl acetals. N -acylglycinal diethyl acetal was dissolved in 25 ml of diethyl ether, 10 ml of saturated NaCl, 7 ml of deionized H2O, and 3 ml of concentrated HCl. Reaction mixture was left stirring for 12 hours, and reaction progress was monitored by TLC (CH2Cl2: EtOAc (80:20)). Organic layer was separated and dried with MgSO4. Solvent was removed under vacuum and the residue was dissolved in CH2Cl2: EtOAc (80:20), and applied to a silica column (5 x 20 cm). N Benzoylglycinal and N -octanoylglycinal were obtain ed in 9 %, and 11 % yields, respectively. Purity and identity of both compounds were confirmed by 1H and 13C NMR spectra obtained on Varian iNova 400 MHz (Figs A-10-11). 13

PAGE 24

Scheme 3. Synthesis of N -Acylglycinals. D. Synthesis of N -Octanoylglycine N -Octanoylglycine was synthesized following the modified procedure of Iyer et al. (40). Glycine (0.02 mol) was dissolved in 100 ml of deionized water, and pH was adjusted to 10 by addition of 2 M NaOH. On e equivalent of octanoyl chloride was added drop-wise to the solution, while mainta ining pH 10 with addition of 2 M NaOH. Following 1.5 hours of reaction time, solution was acidified to pH 1 by addition of 30% H2SO4. White precipitate formed following acidi fication, and was filtered off and dried. Precipitate was then recrystallized with EtOAc /petroleum ether to ob tain the pure product 14

PAGE 25

in 49% yield. N -Octanoylglycine identity and purity was confirm ed by 1H and 13C NMR on Varian iNOVA 400 (Fig A-13). E. Preparation of N -Octanoylglycinal semicarbazone N -Octanoylglycinal semicarbazone st andard was prepared by reacting N octanoylglycinal with a large excess (~10 fo ld) of semicarbazide in aqueous solution. N Octanoylglycinal semicarbazone was purif ied using HPLC and characterized by 1H and 13C NMR on Varian iNova 400 (Fig A-12). Kinetic Assays of NAE Substrate Activity with ADH3 A. Spectrophotometric NADH Assay Kinetic constants for short and mid-le ngth NAEs were determined from initial reaction rates obtained by following NADH production at 340 nm. Reaction conditions were as follows: 100 mM glycine pH 10, 2.5 mM NAD+, and 15-30 g/ml ADH3 in 1 ml total reaction volume. NAE con centrations were between 0.1KM and 10KM. Enzyme assays were conducted at 37 oC, using JASCO V-530 UVVis spectrophotometer. Glycine was purchased from J.T. Baker, sodium pyrophosphate was obtained from Fisher, and NAD+ was acquired from Bioworld. A ll were used without further purification. Rates were normalized using rate for 250 M cinnamyl alcohol as the standard. B. MTS-Formazan Assay Due to the low solubility of long chain NAEs full kinetic assays to determine their kinetic constants could not be performed. As the compounds could only be assayed at relatively low concentrations (<100 M), the rates of NADH production were very low and could not be confidently determined using the spectrophotometric NADH assay. Instead, a more sensitive method for NAD H detection was employed, coupling the NADH production to the reduction of a tetrazo lium dye MTS ([3-(4,5dimethylthiazol-2yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazo lium, inner salt) to the 15

PAGE 26

colored form azan with max at 490 nm and = 20,800 M-1cm-1 at pH 9.5 (Fig. 7). PMS (phenazinemethosulfate) was used as an intermediate electron carrier. Figure 7. MTS-formazan assay for NADH detection. NAEs with 10 or more carbon atoms in the chain were all assayed for ADH3 activity at the same concentration employing the MTS-formazan assay. The assay conditions were as follows: 100 mM s odium pyrophosphate pH 9.5, 55 M NAE, 2.5 mM NAD+, 150 M MTS, 8.25 M PMS, 3 % DMSO and 30 g/ml ADH3. Decanoyl and lauroylethanolamine were soluble enough to allow the determination of KM and VMax values as well. All reac tions were done at 37 oC and the initial rates were determined by observing the increase in absorbance at 490 nm using JASCO V-530 UV-Vis spectrophotometer. MTS was purchased from Promega and PMS was obtained from TCI America. Rates for cinnamyl alcohol obtained in this manner matched the rates obtained by following NADH production at 340 nm. Rates we re normalized using rate for 250 M cinnamyl alcohol as the standard. Modeling Experiments The crystal structure of hum an ADH3 (P DB ID 1MP0 (41)) was used for gridbased ligand docking. All co-crystallized ligands deemed superfluous for enzyme function were removed from the crystal stru cture and polar hydroge ns were added using AutoDockTools. Charges were then corrected for the requisite zinc ions and bond orders corrected for the co-substrate, NAD+. The receptor grid was prepared with a grid point 16

PAGE 27

spacing of 0.2 using AutoGrid. The substrates of interest were then prepared using AutoDockTools to define torsions, rotam ers, and polar hydrogens. The ligands were then docked into the active site of ADH using Au toDock 4.0 (42-43). All default settings were utilized with the exception of increas ing the number of energy evaluations from 2.5 x 104 to 2.5 x 107. Product Characterization Experiments A. HPLC Se paration Experiments Experiments were performed on a HP 1100 Agilent, equipped with a 4-channel solvent mixing system, a quaternary pump, and a deuterium lamp UV detector. All separations were accomplished using a Ther mo-Scientific C18 column (4.6 x 250 mm), with temperature regulated at 40 oC. N -Benzoylethanolamine and its derivatives were separated using either a gradient of 50 mM sodium phosphate pH 6.0: CH3CN 90: 10 to 95:5 over 20 minutes, or isocratic mobile phase of 50 mM sodium phosphate pH 6.0: CH3CN (90:10 (v/v)) (Fig. B-1). Compounds were detected by following UV absorbance at 230 nm. Cinnamyl alcohol and its de rivatives were separated using gradient of 50 mM sodium phosphate pH 6.0: CH3CN 75:25 to 40:60 over 15 minutes, and compounds were detected by monitoring absorbance at 265 nm (Fig B-2). N -Octanoylglycinal semicarbazone and semicarbazide were separated using gradient of H2O: CH3CN 70:30 to 40:60 over 20 minutes (Fig B-3). The compounds were detected by monitoring absorbance at 210 nm. B. GC-MS Experiments GC-MS experiments were performed on Shimadzu GC-MS instrument equipped with a DB-5 (0.25 m x 0.25 mm x 30 m) column. Compounds were extracted from reaction mixture using either ethyl ether (cinnamyl alcohol and cinnamyl aldehyde) or CH2Cl2 ( N -benzoylethanolamine and its derivativ es). Reaction mixture was acidified prior to N -benzoylglycine extraction. Solvent was removed and samples were derivatized prior to analysis in order to increase th eir volatility and si gnal strength. Aldehyde 17

PAGE 28

derivatization to PFB-oxim es was done by disso lving the extracted and dried residue into 90 l of CH3CN, adding 10 l of 100 mM PFBHA ( O -(2,3,4,5,6pentafluorobenzyl)hydro xylamine), and heating the solution at 60 oC for 60 minutes (Scheme 4). Scheme 4. PFBHA derivatization reaction. N -Benzoylglycine derivatiza tion was done by dissolving the extracted and dried material in 100 l of BSTFA ( N,O -bis(trimethylsilyl)trifluoroacetamide), purging the solution with N2, and heating it at 90 oC for 15-20 minutes (Scheme 5). Carboxylic group and amide moiety of NAGs can be both be derivatized by this method, as can hydroxyl and amide groups of NAEs. Scheme 5. BSTFA deri vatization reaction. 18

PAGE 29

Derivatized sam ples (5-10 l) were injected into the GC-MS in a splitless manner, with injection temperature at 250 oC. Temperature program was modified from Merkler et al. (44). Oven temperature was raised from 55.0 oC to 150.0 oC, at a rate of 40.0 oC/min, held at 150.0 oC for 3.6 min, then raised to 300 oC at a rate of 10.0 oC/min, and finally held at 300 oC for 1.0 min. Interface temperature between GC and MS was 280 oC, and solvent cut time was 7-9 min. For N -benzoylglycinal dete ction, injection volume was increased to 40 l in order to more confidently detect the TMS-benzoylglycine derivative. Solvent cut time wa s raised to 11.5 min, so as to avoid column overload by the far more concentrated diand mono-TMSN -benzoylethanolamine derivatives eluting at 10.7 and 11.2 min, respectively. Peak iden tity was established by comparison of retention times and mass spectra with those of derivatized sta ndards and library spectra. C. Trapping N -acylglycinals with semicarbazide In order to characterize the reaction products, N -acylglycinals, which are unstable at reaction pH, semicarbazide was empl oyed as an aldehyde-trapping reagent. Semicarbazide was added in la rge excess (9-10 x substrate co ncentration) to the reaction mix containing 100 mM sodium pyrophosphate pH 9.5, 3-5 mM N -benzoylethanolamine, 2.5 mM NAD+, and 0.25-1.0 mg/ml ADH3. Formation of N -benzoylglycinal semicarbazone was followed by HPLC. D. N -Acylglycinal Semicarbazone Characterization by 13C NMR N -Octanoylglycinal semicarbazone was ch aracterized by following the enzymatic oxidation of [1,2-13C]N -octanoylethanolamine in the presence of semicarbazide by 13C NMR. The presence of N -octanoylglycinal semicarbazone was established by observing the reduction in intensity of the 13C labeled signals in the subs trate and the appearance of two 13C carbon signals consis tent with those of N -octanoylglycinal semicarbazone. Peak identity was confirmed by comparison with 13C spectrum of synthesized N octanoylglycinal semicarbazone (Fig. A-12). Reaction conditions were as follows: 50 mM sodium pyrophosphate pH 8.0, 2.5 mM [1,2-13C]N -octanoylethanolamine, 2.5 mM NAD+, 20 mM semicarbazide HCl, 10% D2O, and 1.7 mg/ml ADH3. Reaction mix was incubated at 37 oC, and 13C NMR spectra were taken af ter 24 hour and 48 hour reaction 19

PAGE 30

tim e points. Control samples containing all th e reagents except for the enzyme were handled and analyzed in the same manner. Experiments were performed on a Varian iNova 400 MHz instrument. 20

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Chapter 3 Results and Discussion ADH3 Purification ADH3 was purified from bovine liver to ~ 90 % homogeneity (Fig. 8), and stored at -80 oC for later use in enzymatic assays. Sp ecific activity of purified ADH3 was 0.53 U/mg, which is somewhat lower that the prev iously reported values of ~ 3.0 U/mg (38). Relatively low activity is most likely due to the fact that enzyme was purified from liver that had been stored in the freezer for se veral months, rather than a fresh sample. Molecular weight of ADH3 subunits, calculated based on the Rf values determined from the SDS-PAGE was 38 kDa, which is in good agreement with previously determined values of 40-41 kDa (32). MWL ADH3 ADH3 MWL 250 kDa 150 kDa 100 kDa 50 kDa 75 kDa 37 kDa 25 kDa Figure 8. SDS-PAGE of purified bovine liver ADH3. The outer two lanes contain the Kaleidascope molecular weight ladder (MWL), and the inner lanes contain purified ADH3 21

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Kinetics of ADH3-catalyzed NAE Oxidation A variety of aliphatic NAEs with chain lengths varying from 2 to 20 carbons, as well as several aromatic NAEs were evaluated as substrates for ADH3, in the manner described previously (Fig. 9). All of the te sted NAEs were found to be substrates for ADH3, with KM values decreasing with increase in chain length of the acyl substituent (Table 1). This indicated that increase in substrate size led to ti ghter binding to the enzyme, or rather that small size impaired the ability to bind to the site tightly, which was in agreement with known substrate pref erences of ADH3 (26) and previous crystallographic studie s of the enzyme active site (30-32). VMax followed the same trend, decreasing with increase in substrate size and hydrophobicity. However, decrease in KM was more dramatic, resulting in overall increase in VMax/KM values with increase in substrate chain length (see Table 1). These fi ndings are in agreement with the previous studies, which established the preference of ADH3 for bulky, longer chain alcohol substrates, in contrast with other ADH en zymes (26). While ADH3 could catalyze the oxidation of all tested NAEs, their turnover values were low compared to cinnamyl alcohol, one of the preferred ADH3 s ubstrates (see Tables 1 and 3). Michaelis-Menten plot of ADH3-catalyzed N -hexanoylethanolamine oxidation[ N -Hexanoylethanolamine] (mM) 0 2040 6080100120Rate (mol/min/mg) 0.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4 1.6 VMAX = 1.6 +/0.018 mol/min/mg KM = 9.5 +/0.36 mM Figure 9. A representiative Michaelis-Mente n plot for one of the assayed NAEs. 22

PAGE 33

Table 1. Kinetic constants for shortand medium-chain NAEs. Substrate R N H O OH KM (mM) VMax (mol/min/mg) VMax / KM N-acetylethanolamine R = CH3 (4.5 0.35) x 102 1.9 0.059 0.0041 N-propionylethanolamine R = CH3CH2 79 3.8 1.9 0.026 0.024 N-butyrylethanolamine R = CH3(CH2)2 46 5.4 1.7 0.052 0.038 N-hexanoylethanolamine R = CH3(CH2)4 9.5 0.36 1.6 0.018 0.17 N-octanoylethanolamine R = CH3(CH2)6 5.8 0.38 1.5 0.028 0.26 N-decanoylethanolamine R = CH3(CH2)8 0.32 0.029 0.57 0.024 1.8 N-lauroylethanolamine R = CH3(CH2)10 0.033 0.0061 0.14 0.0088 4.3 Kinetic constants could not be determined for NAEs longer than lauroyl due to very low solubility and low reaction rates. Table 2 shows a comparison of rates for two medium-chain (decanoyl and lauroyl) and se veral long chain NAEs at a single substrate concentration. Rates continue to follow a decreasing trend observed with short and medium chain NAEs, with the exception of myristoylethanolamine which has a significantly lower rate compared to the longer NAEs tested. It is po ssible that this result is related to the unsaturated bonds presen t in other higher leng th NAEs, which may enable tighter binding to the enzyme. Table 2. Relative ADH3 rates for medium and long-chain NAEs Substrate R N H O OH Relative Rates N-decanoylethanolamine R = CH3(CH2)8 1.0 N-lauroylethanolamine R = CH3(CH2)10 0.57 N-myristoylethanolamine R = CH3(CH2)12 0.061 N-oleoylethanolamine R = CH3(CH2)7HC=CH(CH)7 0.28 N-linoleoylethanolamine R = CH3(CH2)7HC=CHCH2HC=CH(CH)4 0.26 N-arachidonoylethanolamine R = CH3(CH2)3HC=CHCH2HC=CHCH2HC=CHCH2HC=CH(CH)4 0.24 Among the NAEs with an aromatic subs tituent, compounds with benzyl group had lower KM values, higher VMax values, and therefore higher turnover values compared to compounds with a phenyl substituent (see Table 3), indicating that the presence of CH2 23

PAGE 34

group following the arom atic ring might have contributed to bette r binding and more efficient catalysis. Table 3. Kinetic constants for aromatic NAEs and cinnamyl alcohol. Substrate R N H O OH KM (mM) VMax (mol/min/mg) VMax / KM N-benzoylethanolamine R = C6H6 5.2 0.52 0.36 0.0098 0.069 N-phenylacetylethanolamine R = C6H6CH2 3.9 0.24 1.2 0.026 0.32 N-benzyloxycarbonylethanolamine R = C6H5CH2O 2.3 0.14 0.81 0.014 0.36 N-(2-Phenoxyacetyl)ethanolamine R = C6H5OCH2 6.3 0.36 0.25 0.0042 0.040 Cinnamyl alcohol C6H6HC=CHCH2OH 0.035 0.0033 4.0 0.058 4.1 102 Docking Experiments NAEs with chain length between 2 and 11 carbons were docked into the human ADH3 crystal structure using AutoDock 4.0 in order to evaluate their relative binding energies (Table 4). Sequence alignment of human and bovine ADH3 using BLAST (4546) shows that they share 94% identity, w ith 98 % of residues conserved, making the docking results obtained with the human enzy me applicable to bovine ADH3 as well. Relative binding energies of docked NAEs we re found to follow the same general trend as KM values, decreasing with increase in chai n length (Table 4), in dicating that tighter binding to the enzyme is at least pa rtly responsible for the observed KM trend. These findings provide further agreement with previo usly determined substrate preference (26) and crystallographic data (30-32). Substrates were docked in the enzyme active site with hydroxyl group coordinating with the catalytic Zn ion (Fig. 7), in the same manner as other ADH3 substrates have been shown to bind (31-32). 24

PAGE 35

Table 4. Relative binding energies and KM values for selected NAEs and cinnamyl alcohol. Binding energies values were calculated using Autodock 4.0, and have a standard error of about 2.5 kcal/mol (43). Substrate KM (mM) Estimated free energy of binding (kcal/mol) N-acetylethanolamine (4.5 0.35) x 102 -3.72 N-propionylethanolamine 79 3.8 -4.14 N-butyrlethanolamine 46 5.4 -4.22 N-hexanoylethanolamine 9.5 0.36 -4.34 N-octanoylethanolamine 5.8 0.38 -4.31 N-decanoylethanolamine 0.32 0.029 -4.34 Nlauroylethanolamine 0.033 0.0061 -4.06 N-benzoylethanolamine 5.2 0.52 -4.58 Cinnamyl alcohol 0.035 0.0033 -4.68 Figure 10. N-Hexanoylethanolamine docked into the active site of human ADH3 (PDB ID 1MP0). Protein backbone is shown in green, Zn ions are gray spheres, and NAD+ and N -hexanoylethanolamine are shown as stick models with CPK-colored atoms. 25

PAGE 36

Product Characterization Characterization of the product of the en zymatic reaction was initially attempted by following the reaction progress by HPLC. HPLC analysis of the ADH3-catalyzed N benzoylethanolamine oxidation revealed the presence of a peak consistent with N benzoylglycine as well as a peak corresponding to N -benzoylglycinal (Fig. 11). N Benzoylglycine and N -benzoylglycinal are the two po ssible products of the enzymatic oxidation of N -benzoylethanolamine. However, ADH3 has not been found to catalyze the dismutation of aldehydes (18). Experiments using N -benzoylglycinal as a substrate for ADH3 indicated that ADH3 is not able to catalyze the oxidation of N -benzoylglycinal, but that N -benzoylglycinal can undergo non-enzymatic dismutation to N -benzoylglycine and N -benzoylethanolamine under the empl oyed reaction conditions (2.5 mM NAD+, pH 9.5-10). HPLC analysis of ADH3-catalyzed cinnamyl alcohol oxidation revealed only peak consistent with cinnamyl aldehyde (Fi g. 12), further sugges ting that the reaction products of NAE oxidation are N -acylglycinals, and that any NAGs observed are produced through non-enzymatic means. N -benzoylethanolamine N -benzo y l g l y cinal N -benzo y l g l y cine NAD+ derivative NAD+/NADH Figure 11. HPLC separation of ADH3-catalyzed N -benzoylethanolamine oxidation. Reaction conditions were as follows: 100 mM sodium pyrophosphate pH 9.5, 2.5 mM NAD+, 6 mM N benzoylethanolamine, and 2.4 mg/ml ADH3. An aliquot was taken for HPLC analysis after 36 hours of reaction time. Peak at 3.918 was determined to be NAD+ derivative formed during prolonged incubation at high pH and was also observed in the control sample containing only NAD+ and buffer. 26

PAGE 37

cinnamyl alcohol NAD+/NADH c innamyl aldehyde Figure 12. HPLC separation of ADH3-catalyzed cinnamyl alcohol oxidation. Reaction conditions were as follows: 100 mM sodium pyrophosphate pH 9.5, 2.5 mM NAD+, 1.75 mM cinnamyl alcohol, and 5 g/ml ADH3. HPLC analysis was performed after 45 minutes of reaction time. Further characterization of the product peaks observed by HPLC was conducted using GC-MS. In order to test the validity of the method, the proce dure was first used to characterize cinnamyl alcohol reaction produc t. Derivatization of cinnamyl alcohol reaction extract with PFBHA gave two peaks detected by GC-MS, whose retention times and fragmentation patterns were consistent with those for two isomers of cinnamyl aldehyde PFB-oxime (Fig. 13), thus confir ming that the reaction product is cinnamyl aldehyde. However, applying the same method to characterize N -benzoylglycinal observed by HPLC proved unsuccessful. Although derivatized standard for N benzoylglycinal-PFB-oxime was successfully characterized using GC-MS (Fig C-3), the compound could not be detected by this method in the reaction extracts. This can be attributed to the very low turnover rates of N -benzoylethanolamine oxidation compared to cinnamyl alcohol oxidation (Table 3), as well as to the tendency of N -benzoylglycinal to undergo oxidation to N -benzoylglycine under the react ion conditions. Derivatizing reaction extract with BSTFA resulted in detection of N,O -di-TMSN -benzoylglycine (Fig. 14), thus confirming that the peak observed by HPLC is N -benzoylglycine. 27

PAGE 38

Cinnamyl aldehyde O -PFB-oximes (Z and E isomers) Figure 13. GC-MS spectra of cinnamyl aldehyde O -PFB-oximes. Cinnamyl aldehyde PFB-oximes were detected through GC-MS analysis of cinnamyl alcohol reaction extract that was treated with PFBHA. The two peaks correspond to the two cinnamyl aldehyde O -PFB-oxime isomers ( E and Z ), which differ in retention times, but have identical fragmentation patterns. See Fig. C-1 for cinnamyl aldehyde O -PFBoxime standard spectra. O N F F F F F 77 91 130 181 (1 E ,2 E )-cinnamaldehyde O -perfluorobenzyloxime ChemicalFormula:C16H10F5NO ExactMass:327.07 28

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N,O -di-TMSN -benzoylglycine Figure 14. GC-MS spectrum of N,O-di-TMSN -benzoylglycine. N,O -di-TMSN -benzoylglycine peak was detected in derivatized extract of ADH3-catalyzed N -benzoylethanolamine reaction. Extraction and GC-MS analysis were performed after 24 hours of reaction time. In order to detect a nd characterize the unstable N -acylglycinal product, a wellknown aldehyde-trapping reagent, semicarbazide, was added to the reaction mixture. Semicarbazide readily reacts with aldehydes and ketones to form semicarbazones (Fig. 15). HPLC analysis of N -benzoylethanolamine reaction containing semicarbazide revealed a peak consistent with the peak for N -benzoylglycinal semicarbazone standard (Figs.12 and B-1). Further characterization of the semicarbazone derivative was attempted by LC-MS and GC-MS, but proved unsuccessful. N O O O Si N,O -di-TMSN -benzoylglycineSi 45 73 105 206 73 308 77 ChemicalFormula:C15H25NO3Si2ExactMass:323.14 29

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O R1 R2 H2N N H NH2 O + N N H NH2 O R1 R2 Aldehyde/Ketone Semicarbazide Semicarbazone Figure 15. Semicarbazone formation reaction. N -benzoylethanolamine NAD+ derivative NAD+/NADH N -benzoylglycinal semicarbazone Figure 16. HPLC analysis of ADH3-catalyzed N -benzoylethanolamine oxidation in the presence of semicarbazide. Reaction conditions were as follows: 100 mM sodium pyrophosphate pH 9.5, 2.5 mM NAD+, 4 mM N -benzoylethanolamine and 300 g/ml ADH3. HPLC analysis was performed after 24 hours of reaction time. In order to characterize the putative N -acylglycinal semicarbazone species detected by HPLC, reaction was analyzed by 13C NMR. For the purpose of increased sensitivity, [1,2-13C]N -octanoylethanolamine was synt hesized and employed as the reaction substrate. Analysis of [1,2-13C]N -octanoylethanolamine reaction showed a total of 4 different 13C signals two that matched those for the substrate, at 41 and 60 ppm (Fig. A-9), and two new 13C signals, at 40 and 143 ppm. Chemical shifts of the two new signals match those of th e corresponding carbon atoms in the N -octanoylglycinal semicarbazone standard (Figs. 17 and A-12), thus providing confirmation that the product of the ADH3 catalyzed oxidation is N -acylglycinal, in agreement with previously published data (18). Performing the reaction in absence of semicarbazide resulted in 30

PAGE 41

appearan ce of 13C signals consistent with those of N -octanoylglycine (Figs. 18 and A-13). This result was consistent with previous HPLC experiments, which showed that N benzoylglycinal is readily oxidized to N -benzoylglycine at basic pH. Control sample with no ADH3 present showed only the two substrat e peaks (Fig A-14), confirming there was no detectable oxidation of N -octanoylethanolamine through non-enzymatic means. B A A2 B2 Figure 17. 13C NMR spectrum of ADH3-catalyzed [1,2-13C]-N -octanoylethanolamine oxidation in the presence of semicarbazide. Reaction conditions were as indicated in Chapter 2, and spectrum was taken after 48 hours of reaction time. 31

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A B A3 B3 Figure 18. 13C NMR spectrum of ADH3-catalyzed [1,2-13C]-N -octanoylethanolamine oxidation without semicarbazide. Reaction conditions were as indicated in Chapter 2 with the exception of semicarbazide, which was omitted. Spectrum wa s taken after 48 hours of reaction time. Conclusion Evaluation of NAEs as ADH3 substrates s howed that they can be oxidized by this enzyme, which is known to act on a variety of midand long-chain alcohol substrates. In accordance with previous literature reports, the reaction products were shown to be N acylglycinals, which ADH3 was unable to oxidize further to NAGs. Relatively low conversion rates observed for ADH3-cataly zed NAE oxidation make it unlikely that ADH3 plays a significant role in conversion of NAEs to N -acylglycinals in vivo Nevertheless, our findings validate the chem istry proposed in the pathway, and further suggest that another ADH enzyme could be re sponsible for the oxidative degradation of NAEs. Further studies on ADH5 and ADH6 should be completed in order to evaluate the possibility of their involvement in NAE metabol ism, as these two enzymes have as of yet been poorly investigated and could potentiall y play the role of FADH in the conversion 32

PAGE 43

of NAEs to NAGs. Additionally, NAEs could be ev aluated as substrates or inhibitors of other ADH enzym es, in order to explore thei r potential role in alcohol or retinol metabolism. 33

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References Cited 1. Farrell, E. K., and Merkler, D. J. (2008) Biosynthesis, degradation and pharm acological importance of fatty acid amides. Drug Discov. Today. 13, 558-568. 2. Schmid, H.H.O., and Berdyshev E. V ., (2002) Cannabinoid receptor-inactive N acylethanolamines and other fatty acid amides: metabolism and function. Prostaglandins, Leukot. Essent. Fatty Acids. 66, 363-376. 3. Lambert, D. M., and Muccioli, G. G. (2007) Endocannabinoids and related N acylethanolamines in the control of appetite and energy metabolism: emergence of new molecular players. Curr. Opin. Nutr. Metab. Care. 10, 735-744. 4. Hansen, H. S., Moesgaard, B., Petersen G., and Hansen, H. H. (2002) Putative neuroprotective actions of N -acylethanolamines. Pharmacol. Ther. 95, 119-126. 5. Lambert, D. M., Vandervoorde, S., Jon sson, K., and Fowler, C. J. (2002) The palmitoylethanolamide family: a new class of anti-inflammatory agents? Curr. Med. Chem. 9, 663-674. 6. Dalle Carbonare, M., Del Giudice, E., Stecca, A., Colavito, D., Fabris, M., DArrigo, A., Bernardini, D., Dam, M., and Leon, A. (2008) A saturated N -acylethanolamine other than N -palmitoyl ethanolamine with anti-i nflammatory properties: a neglected story. J. Neuroendocrinol. 20 (suppl.1), 26-34. 7. Jonsson, K., Vandervoorde, S., Lambert, D. M., Tiger, G., and Fowler, C. J. (2001). Effects of homologues and analogues of palmityolethanolamide upon the activation of the endocannabinoid anandamide. Br. J. Pharmacol. 133, 1263-1275. 8. Lambert, D. M., DiPaolo, F. G., S onveaux, P., Kanyonyo, M., Govaerts, S. J., Hermans, E., Bueb, J., Delzenne, N. M., and Tschirhart, E. J. (1999) Analogues and homologues of N -palmitoylethanolamide, a putative endogenous CB2 cannabinoid, as potential ligands for the cannabinoid receptors. Biochim. Biophys. Acta 1440, 266274. 34

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9. Tsuboi, K., Takezaki, N ., and Ueda, N. (2007) The N -acylethanolamine-hydrolyzing acid amidase (NAAA). Chem. Biodivers 4, 1914-1925. 10. Ueda, N., Tsuboi, K., and Lambert, D. M. (2005) A second N -acylethanolamine hydrolase in mammalian tissues. Neuropharmacology 48, 1079-1085. 11. Arafat, E. S., Trimble, J. W., Andersen, R. N., Dass, C., and Desiderio, D. M. (1989) Identification of fatty acid amides in human plasma. Life Sci 45, 1679-1687. 12. Burstein, S. H., Rossetti, R. G., Yagen, B., and Zurier R. B. (2000) Oxidative metabolism of anandamide. Prostaglandins & other Lipid Mediat. 61, 29-41. 13. Sugiura, K., Kondo, S., Kodaka, T., Tonegawa, T., Nakane, S., Yamashita, A., Ishima, Y., and Waku, K. (1996). Enzyma tic synthesis of oleamide (cis-9, 10octadecenoamide), and endogenous sleep-i nducing lipid, by rat brain microsomes. Biochem. Mol. Biol. Int 40, 931-938. 14. Mueller, G. P., and Driscoll, W. J. (2007) In vitro synthesis of oleoylglycine by cytochrome c points to a novel pathway for the production of lipid signaling molecules. J. Biol. Chem. 282 22364-22369. 15. Driscoll, W. J., Chaturvedi, S., and Mueller, G. P. (2007) Oleamide synthesizing activity from rat kidney identification as cytochrome c. J. Biol. Chem. 282 2235322363. 16. Jrnvall, H. and Hg, J. (1995) Nome nclature of alcohol dehydrogenase. Alcohol Alcohol. 30, 153-161 17. Yin, S., Han, C., Lee, A., and Wu, C. (1999) Human alcohol dehydrogenase family. Adv. Exp. Med. Biol. 463, 265-274. 18. Svensson, S., Lundsj, A., Cronholm, T., and Hg, J. (1996) Aldehyde dismutase activity of human liver alcohol dehydrogenase. FEBS Lett. 394, 217-220. 19. Yin, S., Chou, C., Lai, C., Lee, S., a nd Han, C. (2003) Human class IV alcohol dehydrogenase: kinetic mechanism, func tional roles and medical relevance. Chem. Biol. Interact. 143-144, 219-227. 20. Hg, J., Hedberg, J. J., Strmberg, P., a nd Svensson, S. (2001) Mammalian alcohol dehydrogenases functional and structural implications. J Biomed Sci. 8 71-76. 21. Duester, G. (2000) Fam ilies of retinoid dehydrogenases vitamin A function production of visual pigment and retinoic acid. Eur. J. Biochem. 267 4315-4324. 35

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22. Jrnvall, H., Hg, J., Persson B., and Pa rs, X. (2000) Pharm acogenetics of the alcohol dehydrogenase system. Pharmacology. 61, 184-191. 23. Yasunami, M., Chen, C., and Yoshida, A. (1999) A human alcohol dehydrogenase gene (ADH6) encoding an additional class of isozyme. Proc. Natl. Acad. Sci. 88, 7916-7614. 24. Zheng, Y., Bey, M., Liu, H., and Felder, M. R. (1993) Molecular basis of the alcohol dehydrogenase-negative deer mouse. J. Biol. Chem. 268, 24933-24939. 25. Pars, X., and Vallee, B. L. (1981) Ne w human liver alcohol dehydrogenase forms with unique characteristics. Biochem. Biophys. Res. Commun 98, 122-130. 26. Wagner, F. W., Pars, X., Hol mquist, B., and Vallee, B. L. (1984) Physical and Enzymatic Properties of a Class I II Isozyme of Human Liver Alcohol Dehydrogenase: -ADH. Biochemistry, 23, 2193-2199. 27. Koivusalo, M., Baumann, M., and Uotila, L. (1989) Evidence for the identity of glutathione-dependent formaldehyde dehydrogenase and class III alcohol dehydrogenase. FEBS Lett. 257, 105-9. 28. Holmquist, B., and Vallee, B. L. (1991) Human liver class III alc ohol and glutathione dependent formaldehyde dehydrogenase are the same enzyme. Biochem. Biophy.s Res. Commun. 178, 1371-1377. 29. Lee, S., Wang, M., Lee, A., and Yin, S. ( 2003) The metabolic role of human ADH3 functioning as ethanol dehydrogenase. FEBS Lett. 544 143-147. 30. Yang, Z., Bosron, W. F., and Hurley, T. D., (1997) Structure of human alcohol dehydrogenase: a glutathione-depende nt formaldehyde dehydrogenase. J. Mol. Biol. 265, 330-343. 31. Sanghani, P. C., Bosron, W. F., and Hurl ey, T. D. (2002) Human glutathionedependent formaldehyde dehydrogenase. Struct ural changes associated with ternary complex formation. Biochemistry. 41, 15189-15194. 32. Sanghani, P. C., Robinson, H., Bosron, W. F., and Hurley, T. D. (2002) Human glutathione-dependent formaldehyde dehydroge nase. Structures of apo, binary, and inhibitory ternary complexes. Biochemistry. 41, 10778-10786. 36

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33. Ra maswamy, S., Eklund, H., and Plapp, B. V. (1994). Structures of horse liver alcohol dehydrogenase complexed with NAD+ and substituted benzyl alcohols. Biochemistry, 33, 5230-5237. 34. Eklund, H., and Brndn, C. (1979). Stru ctural differences between apoand holoenzyme of horse liver alcohol dehydrogenase. J. Biol. Chem. 254, 3458-3461. 35. Colonna-Cesari, F., Perahia, D., Karplus, M., Eklund, H., Brndn, C. I., and Tapia, O. (1986) Interdomain motion in liver alcohol dehydrogenase: structural and energetic analysis of the hinge bending mode. J. Biol. Chem. 261, 15273-15280. 36. Sanghani, P. C., Stone C. L., Ray, B. D., Pi ndel, E. P., Hurley, T. D, and Bosron, W. F. (2000) Kinetic mechanism of human glutathione-dependent formaldehyde dehydrogenase. Biochemistry. 39, 10720-10729. 37. Giri, P. R., Linnoila, M., ONeill, J. B ., and Goldman, D. (1989) Distribution and possible metabolic role of class III alcohol dehydrogenase in the human brain. Brain Res. 481, 131-141. 38. Pourmotabbed, T., and Creighton, D. J. (1986) Substrate specificity of bovine liver formaldehyde dehydrogenase. J.Biol. Chem. 261 14240-14244. 39. Brown, E. V. (1949) Penniloaldehydes and penaldic acids. In Chemistry of penicillin ; Clarke, H. T., Johnson, J. R., and Robinson, R., Eds. Princeton University Press: Princeton, NJ. p 482-483. 40. Iyer, V. I., Sheth, G. N., and Subrahmanyam, V. V. R. (1982) Melting behaviour of some pure N -acyl amino acids and peptides. J. Indian Chem. Soc. 60, 856-859. 41. Sanghani, P. C., Robinson, H., Bennet-Lovsey, R., Hurley, T. D., and Bosron, W. F. (2003) Structure-function relationships in human class III alcohol dehydrogenase (formaldehyde dehydrogenase). Chem. Biol. Interact.. 143-144, 195-200. 42. Morris, G. M., Goodsell, D. S., Halliday, R.S., Huey, R., Hart, W. E., Belew, R. K. and Olson, A. J. (1998) Automated docki ng using a Lamarckian genetic algorithm and empirical binding free energy function. J. Comput. Chem. 19, 1639-1662. 43. Huey, R., Morris, G. M., Olson, A. J. and Goodsell, D. S. (2007) A semiempirical free energy force field with charge-based desolvation. J. Comput. Chem. 28, 11451152. 37

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44. Merkler, D. J., Chew, G. H., Gee, A. J., Merkler, K. A., Sorondo, J. O., and Johnson, M. E. Oleic acid derived m etabol ites in mouse neuroblastoma N18TG2 cells. Biochemistry. 43, 12667-12674. 45. Altschul, S. F., Madden, T. L., Schffer, A. A., Zhang, J., Zhang, Z., Miller, W., and Lipman, D. J. (1997) Gapped BLAST and PS I-BLAST: a new generation of protein database search programs. Nucleic Acids Res. 2, 3389-3402. 46. Altschul, S. F., Wootton, J. C., Gertz, E. M., Agarwala, R., Morgulis, A., Schffer, A. A., and Yu, Y. (2005) Protein database searches using compositionally adjusted substitution matrices. FEBS J. 272, 5101-5109. 38

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APPENDICES 39

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Appendix A NMR Spe ctra Figure A-1. 1H and 13C NMR Spectra of N -Butyrylethanolamine in DMSO-d6. Peak at 3.409 ppm in the 1H spectrum is due to presence of a small amount of H2O in the solvent. 40

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Appendix A (Continued) N H OH O N -Hexanoylethanolamine Figure A-2. 1H and 13C NMR Spectra of N -Hexanoylethanolamine in DMSO-d6. 41

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Appendix A (Continued) N H OH O N -Octanoylethanolamine Figure A-3. 1H and 13C NMR Spectra of N -Octanoylethanolamine in DMSO-d6. 42

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Appendix A (Continued) Fi gure A-4. 1H and 13C NMR Spectra of N -Decanoylethanolamine in DMSO-d6. 43

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Appendix A (Continued) Fi gure A-5. 1H and 13C NMR Spectra of N -Lauroylethanola mine in DMSO-d6. 44

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Appendix A (Continued) Figure A-6. 1H and 13C NMR Spectra of N -Myristoylethanolamine in DMSO-d6. Peak at 3.280 ppm in the 1H spectrum is due to presence of a small amount of H2O in the solvent. 45

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Appendix A (Continued) Figure A-7. 1H and 13C NMR Spectra of N -Oleoylethanolamine in DMSO-d6. 46

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Appendix A (Continued) Fi gure A-8. 1H and 13C NMR Spectra of N -Benzoylethanola mine in DMSO-d6. Peak at 3.398 ppm in the 1H spectrum is due to presence of a small amount of H2O in the solvent. 47

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Appendix A (Continued) Figure A-9. 1H and 13C NMR Spectra of [1,2-13C]-N -Octanoylethanolamine in DMSO-d6. Peak at 3.287 ppm in the 1H spectrum is due to presence of a small amount of H2O in the solvent. 13C-labeled atoms are denoted by *. 48

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Appendix A (Continued) Figure A-10. 1H and 13C Spectra of N -Benzoylglycinal in CDCl3. Peak at 1.598 ppm in the 1H spectrum is due to presence of a small amount of H2O in the solvent. 49

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Appendix A (Continued) Figure A-11. 1H and 13C NMR Spectra of N -Octanoylglycinal in CDCl3. 50

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Appendix A (Continued) Figure A-12. 1H and 13C NMR Spectra of N -Octanoylglycinal semicarbazone in DMSO-d6. Peak at 1.598 ppm in the 1H spectrum is due to presence of a small amount of H2O in the solvent. Signal for C atom marked by is obscured by the DMSO solvent peak in the 13C spectrum. 51

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Appendix A (Continued) N H OH O O N -Octanoylglycine Figure A-13. 1H and 13C NMR Spectra of N -Octanoylglycine in DMSO-d6. 52

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Appendix A (Continued) Figure A-14 13C NMR spectrum of [1,2-13C]N -octanoylethanolamine co ntrol sample without ADH3. 53

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Appendix B HPLC Spe ctra (a) (b) N -benz oylglycine OFigure B-1. HPLC separation for N -benzoylethanolamine and its derivatives. (a) Mobile phase: gradient of 90:10 (50 mM sodium phosphate pH 6.0: CH3CN) to 95:5 over 20 minutes. (b) Mobile phase: 90:10 (50 mM sodium phosphate pH 6.0: CH3CN). N -benzoylethanolamine N -benzoylglycinal N -benzoylglycinal semicarbaz one N H OH O O ON H O OH N H N N H N -benzoylethanolamine N -benzoylglycinal N -benzoylglycine NH2 54

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Appendix B (Continued) Figure B-2. HPLC separation of cinnamyl alcohol and its derivatives. Figure B-3. HPLC separation of semicarbazide and N -octanoylglycinal semicarbazone. S e micarb az id e N -Octanoylglycinal semicarbazone Cinnamy l a lcohol Cinnamyl aldehyde O OH Cinnamic a c i d 55

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Appendix C GC-MS Spectra Cinnamyl aldehyde O -PFB-oximes (E and Z isomers) PFBHA Figure C-1. GC-MS Spectra of cinnamyl aldehyde O -PFB-oxime standard. O N F F F F F 77 91 130 181 (1 E ,2 E )-cinnamaldehyde O -perfluorobenzyloxime ChemicalFormula:C16H10F5NO ExactMass:327.07 56

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Appendix C (Continued) N,O-di-TMS N -benzoylglycine N -TMS N -benzoylglycine Figure C-2. GC-MS spectra of TMS-derivatized N -benzoylglycine standard. N O O O Si N,O -di-TMSN -benzoylglycineSi 73 105 206 73 308 77 ChemicalFormula:C15H25NO3Si2ExactMass:323.14 N O OH O Si 51 73 105 206 N -TMSN -benzoylglycine 77 Ch emicalFormula:C12H17NO3Si ExactMass:251.10 236 57

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Appendix C (Continued) N -benzoylglycinalO -PFB-oxime Figure C-3. GC-MS Spectrum of N -benzoylglycinal O -PFB-oxime standard. E and Z isomers are not resolved. 58


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N-acylethanolamines as novel alcohol dehydrogenase 3 substrates
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ABSTRACT: N-Acylethanolamines (NAEs) are a class of fatty acid amides that act as important mammalian signaling molecules. N-Arachidonoylethanolamine is the best-studied representative and is one of the endogenous ligands for endocannabinoid receptors. NAEs play a role in the regulation of appetite, act as anti-inflammatory and analgesic agents, and are thought to have a neuroprotective function as well. They have been proposed to also serve as precursors to N-acylglycines (NAGs). N-Acylglycinals are likely to be intermediates between the NAEs and the NAGs. The sequential actions of a putative fatty alcohol dehydrogenase and a putative fatty aldehyde dehydrogenase are thought to affect the NAD+-dependent oxidation of the NAEs to the NAGs. NAGs, in turn, serve as precursor in the biosynthesis of primary fatty acid amides (PFAMs), another class of mammalian regulatory molecules. Alcohol dehydrogenase 3 (ADH3), an enzyme known to oxidize mid- and long-chain alcohols to aldehydes, was evaluated for its potential in oxidation of NAEs to N-acylglycinals. In order to evaluate the possibility of ADH3 involvement in NAE metabolism, variable chain length NAEs were synthesized and evaluated as substrates for bovine liver ADH3. NAEs were oxidized by ADH3 in the presence of NAD+, yielding the corresponding N-acylglycinals. V[subfield Max]/K[subfield M] values for assayed NAEs were low relative to cinnamyl alcohol, one of the preferred substrates for ADH3. Our data suggest that the ADH-mediated oxidation of NAEs could occur in vivo, but that ADH3 is unlikely to be the in vivo catalyst.
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