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Keywords:

  • 2-arachidonoylglycerol;
  • anandamide;
  • HRASLS family;
  • N-acyl-phosphatidylethanolamine;
  • phospholipid

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References

Endocannabinoids are endogenous ligands of the cannabinoid receptors CB1 and CB2. Two arachidonic acid derivatives, arachidonoylethanolamide (anandamide) and 2-arachidonoylglycerol, are considered to be physiologically important endocannabinoids. In the known metabolic pathway in mammals, anandamide and other bioactive N-acylethanolamines, such as palmitoylethanolamide and oleoylethanolamide, are biosynthesized from glycerophospholipids by a combination of Ca2+-dependent N-acyltransferase and N-acyl-phosphatidylethanolamine-hydrolyzing phospholipase D, and are degraded by fatty acid amide hydrolase. However, recent studies have shown the involvement of other enzymes and pathways, which include the members of the tumor suppressor HRASLS family (the phospholipase A/acyltransferase family) functioning as Ca2+-independent N-acyltransferases, N-acyl-phosphatidylethanolamine-hydrolyzing phospholipaseD-independent multistep pathways via N-acylated lysophospholipid, and N-acylethanolamine-hydrolyzing acid amidase, a lysosomal enzyme that preferentially hydrolyzes palmitoylethanolamide. Although their physiological significance is poorly understood, these new enzymes/pathways may serve as novel targets for the development of therapeutic drugs. For example, selective N-acylethanolamine-hydrolyzing acid amidase inhibitors are expected to be new anti-inflammatory and analgesic drugs. In this minireview, we focus on advances in the understanding of these enzymes/pathways. In addition, recent findings on 2-arachidonoylglycerol metabolism are described.


Abbreviations
ABHD

α/β-hydrolase domain-containing protein

2-AG

2-arachidonoylglycerol

COX

cyclooxygenase

DAG

diacylglycerol

DAGL

diacylglycerol lipase

FAAH

fatty acid amide hydrolase

FLAT

fatty acid amide hydrolase-like anandamide transporter

GDE1

glycerophosphodiester phosphodiesterase 1

GP-NAE

glycerophospho-N-acylethanolamine

HEK

human embryonic kidney

HRASLS

HRAS-like suppressor

LPA

lysophosphatidic acid

LPS

lipopolysaccharide

LRAT

lecithin retinol acyltransferase

MAG

monoacylglycerol

MAGL

monoacylglycerol lipase

mGluR

metabotropic glutamate receptor

NAAA

N-acylethanolamine-hydrolyzing acid amidase

NAE

N-acylethanolamine

NAPE

N-acyl-phosphatidylethanolamine

NAPE-PLD

N-acyl-phosphatidylethanolamine-hydrolyzing phospholipase D

OEA

oleoylethanolamide

PA

phosphatidic acid

PEA

palmitoylethanolamide

PHARC

polyneuropathy, hearing loss, ataxia, retinitis pigmentosa, and cataract

PL

phospholipase

PLA/AT

phospholipase A/acyltransferase

pNAPE

N-acyl-plasmenylethanolamine

PtdCho

phosphatidylcholine

PtdEtn

phosphatidylethanolamine

PtdIns

phosphatidylinositol

(S)-OOPP

(S)-N-(2-oxo-3-oxetanyl)-3-phenylpropionamide

Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References

Endocannabinoids are endogenous ligands of the cannabinoid receptors CB1 and CB2 [1, 2]. Arachidonoylethanolamide (anandamide) [3] and 2-arachidonoylglycerol (2-AG) [4, 5] are considered to be two physiologically important endocannabinoids (Fig. 1). Although both lipid molecules possess an arachidonic acid chain in their structure, the former is an N-acylethanolamine (NAE), and the latter is a monoacylglycerol (MAG). It should be noted that anandamide is a minor component in animal tissues, whereas other NAEs, such as palmitoylethanolamide (PEA), stearoylethanolamide, oleoylethanolamide (OEA), and linoleoylethanolamide, are abundant [6]. PEA exerts anti-inflammatory, analgesic and neuroprotective effects, and functions as an agonist of peroxisome proliferator-activated receptor-α rather than CB1 or CB2 [7]. OEA is noted for its appetite-suppressing effect, and has been reported to be an agonist of not only peroxisome proliferator-activated receptor-α, but also of transient receptor potential vanilloid 1 and the G protein-coupled receptor GPR119 [8]. By analogy, MAGs other than 2-AG may also show cannabinoid receptor-independent activities. In fact, like OEA, 2-oleoylglycerol activates GPR119 and stimulates the release of glucagon-like peptide-1 in vivo [9].

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Figure 1. Chemical structures of endocannabinoids and related compounds. 2-OG, 2-oleoylglycerol.

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It is believed that, in response to a variety of cellular stimuli, endocannabinoids and related bioactive lipid molecules are principally generated from membrane phospholipids by specific hydrolases or a combination of acyltransferases and hydrolases. After functioning as receptor ligands, these molecules are degraded by hydrolases. Generally, the synthesizing enzymes for bioactive lipid molecules are tightly regulated and expressed at much lower levels than the degrading enzymes, enabling their generation on demand and quick degradation. Despite their structural similarities, the metabolic pathways of anandamide (Fig. 2) and 2-AG (Fig. 3) in animal tissues are completely different [10-12]. The ‘canonical pathways’ for both endocannabinoids are relatively simple. However, recent studies, including the analysis of gene-deficient mice, have shown that: (a) the pathways are more complicated than expected; and (b) one reaction is often catalyzed by multiple enzymes or isozymes. As specific inhibitors of ‘endocannabinoid hydrolases’, such as fatty acid amide hydrolase (FAAH), MAG lipase (MAGL), and NAE-hydrolyzing acid amidase (NAAA), as well as other related enzymes, are expected to be therapeutic drugs [13], it is crucial to elucidate the complete pathways involved in endocannabinoid metabolism in humans and rodents. In the present article, we will briefly discuss the metabolic pathways for anandamide and other NAEs, with special reference to the ‘alternative pathways’. We will also review the new findings on 2-AG metabolism. Although we focus on mammals, it should be noted that some of these pathways are conserved in other organisms, such as frogs, fishes, worms, Arabidopsis, and Dictyostelium, and play unique roles [14-18].

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Figure 2. Metabolic pathways of anandamide and other NAEs. Asterisks indicate the enzymes included in the canonical pathway. Ca-NAT, Ca2+-dependent N-acyltransferase; PTPN22, protein tyrosine phosphatase, nonreceptor type 22; SHIP1, Src homology 2 domain-containing inositol-5-phosphatase 1; sPLA2, secretory PLA2.

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Figure 3. Metabolic pathways of 2-AG.

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Canonical pathways for anandamide and other NAEs

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References

Schmid and his group delineated the metabolic pathways of NAE on the basis of their biochemical studies with animal tissues such as heart and brain [19]. The classic biosynthetic route, composed of two enzyme reactions, was called ‘the transacylation–phosphodiesterase pathway’. The intermediate metabolite, N-acylated ethanolamine phospholipid, is an unusual glycerophospholipid molecule in which an extra acyl chain is covalently linked to the amino group of ethanolamine phospholipid [10, 20, 21]. N-acylated ethanolamine phospholipid species comprise the diacyl type [N-acyl-phosphatidylethanolamine (NAPE)], the alkenylacyl type [plasmalogen-type, N-acyl-plasmenylethanolamine (pNAPE)], and the alkylacyl type (N-acyl-plasmanylethanolamine), which have an acyl chain, an alk-1-enyl chain, or an alkyl chain, respectively, at the sn-1 position (Fig. 4). Alternatively, NAPE may be used as a generic term for all three types of N-acylated ethanolamine phospholipid. In addition to serving as a precursor of various NAEs, NAPE seems to function in membrane stabilization [22]. NAPE was also reported to be an anorexic hormone [23]. However, this finding has been questioned, because a decrease in food intake was also observed after intraperitoneal injection of the negative control phosphatidylethanolamine (PtdEtn) into mice [24]. Regarding the degradation of NAEs, hydrolysis to free fatty acid and ethanolamine is a major pathway [25, 26]. Oxygenation of the arachidonic acid moiety of anandamide appears to be another degradative pathway [27, 28].

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Figure 4. Chemical structures of N-acylated ethanolamine phospholipids.

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Ca2+-dependent N-acyltransferase

The first reaction of NAE biosynthesis is N-acylation of diacyl-type (PtdEtn), alkenylacyl-type (plasmalogen) and alkylacyl-type ethanolamine phospholipids, leading to the formation of N-acylated ethanolamine phospholipids such as NAPE (Fig. 2). The enzyme responsible, ‘N-acyltransferase’, has several unique properties [29-31]. First, the enzyme uses phosphatidylcholine (PtdCho), 1-acyl-lyso-PtdCho, PtdEtn and cardiolipin as donor substrates, and extracts an acyl group selectively from the sn-1 position of these donor phospholipids. Thus, 2-acyl-lysophospholipid, a lysophospholipid that is typically produced by phospholipase (PL)A1, is another product in this reaction. Acyl-CoA is not utilized, although this molecule serves as a donor substrate for many other acyltransferases. Interestingly, NAPE synthase in plants catalyzes direct incorporation of free fatty acid into PtdEtn [18]. Second, the enzyme is associated with membranes, and can be solubilized with the nonionic detergent Nonidet P-40 [31]. Third, the activity of the enzyme is stimulated by Ca2+. It is well known that NAPE accumulates in ischemic heart and brain, or after toxic insult to tissues or cells [29, 32]. Recently, Janfelt et al. showed, with desorption ESI imaging MS, that, during ischemia–reperfusion in the brain of neonatal rats, the levels of many NAPE species increased in the whole injured area, where the cells seemed to be dead [33]. These findings suggest that a remarkable increase in intracellular Ca2+ levels activates N-acyltransferase to generate NAPE. However, it is unclear whether a modest increase in Ca2+ levels caused by physiological stimuli potentiates the enzymatic activity. Despite efforts over many years this Ca2+-dependent N-acyltransferase remains molecularly uncharacterized.

NAPE-hydrolyzing PLD (NAPE-PLD)

The second step in the canonical pathway is the release of NAE from NAPE by a PLD-type enzyme known as NAPE-PLD [34]. cDNA cloning revealed that NAPE-PLD belongs to the metallo-β-lactamase family and is molecularly distinguished from the known PLD isoforms, which hydrolyze common glycerophospholipids such as PtdCho to produce phosphatidic acid (PA), an intracellular signal molecule [35]. NAPE-PLD appears to contain catalytically essential zinc, and the purified recombinant enzyme is specific for NAPE, being almost inactive with major glycerophospholipids such as PtdCho and PtdEtn [36]. At this time, NAPE-PLD is the sole enzyme in animal tissues known to directly release NAE from NAPE. This is in marked contrast to plants, which lack NAPE-PLD, but express multiple PLD isoforms, such as PLDβ and PLDγ, that hydrolyze different glycerophospholipids, including NAPE [16]. NAPE-PLD is tightly bound to membranes. The soluble enzyme prepared by treatment with detergent can be stimulated by millimolar concentrations of divalent cations, including Ca2+ and Mg2+ [37], and 10–100 μm PtdEtn [38]. However, the physiological regulators of NAPE-PLD activity are unknown. Concerning transcriptional regulation, lipopolysaccharide (LPS) downregulates the expression of NAPE-PLD mRNA in RAW264.7 macrophage cells [39]. LPS altered the acetylation state of histone proteins bound to the NAPE-PLD promoter and suppressed the transcription of NAPE-PLD mRNA [40]. The transcription factor Sp1 was involved in the regulation of baseline NAPE-PLD expression, but not in the suppression by LPS. In addition, the expression of NAPE-PLD in rodent brains is age-dependently upregulated at the mRNA and protein levels [41], in agreement with the increase in NAPE-PLD activity [41, 42]. In contrast, N-acyltransferase activity decreases during development [42]. The opposite changes of these two enzymes probably explain why the NAPE level in the ischemic brain of young rodents is much higher than that in adult rodents.

NAPE-PLD−/− mice were born at the expected Mendelian frequency, were viable, and were apparently healthy [43, 44]. The accumulation of NAPEs in the brains of NAPE-PLD−/− mice demonstrated an important role of NAPE-PLD in the degradation of NAPE in this tissue. However, the detection of unaltered or moderately reduced NAE levels strongly suggested the existence of other NAE-forming enzyme(s) or route(s) in NAPE-PLD−/− mice. Apart from this gene-modified mouse model, a recent cohort study suggested a physiological role of NAPE-PLD. In this study, a common haplotype in NAPE-PLD was reported to be protective against severe obesity in a Norwegian population [45].

NAEs with different N-acyl chains appear to share the transacylation–phosphodiesterase pathway for their biosynthesis. In fact, N-acyl species were not distinguished in the reactions catalyzed by N-acyltransferase [46] and NAPE-PLD [36]. The N-acyl chain originates from the sn-1 position of the donor phospholipid, and, in general, polyunsaturated fatty acyl chains such as the arachidonoyl chain are mostly linked to the sn-2 rather than the sn-1 position. Most likely, this is the main reason why the tissue levels of anandamide are usually much lower than those of saturated or monounsaturated NAEs. Apart from this pathway, anandamide may be formed by the condensation of free arachidonic acid with ethanolamine in the reverse reaction of FAAH (Fig. 2) [47, 48]. This reverse reaction can occur if both arachidonic acid and ethanolamine are present in sufficient amounts [49]. Anandamide was actually formed through this route in vivo after partial hepatectomy of mice [50]. Moreover, anandamide could be spontaneously formed from arachidonoyl-CoA and ethanolamine [51].

FAAH

A membrane-associated amidohydrolase that hydrolyzes anandamide and other NAEs to their corresponding fatty acids and ethanolamine has been extensively studied [25, 26]. When its cDNA cloning was reported, the enzyme was named FAAH [52]. FAAH is a serine hydrolase belonging to the amidase signature family. The catalytic triad is composed of Lys142, Ser217, and Ser241 (the catalytic nucleophile) [53]. Analysis of FAAH−/− mice demonstrated the central role of FAAH in the degradation of not only anandamide but also other NAEs, including PEA and OEA [54, 55]. FAAH also hydrolyzes other fatty acid amides, such as oleamide (primary amide of oleic acid) [52] and N-acyltaurine (taurine-conjugated fatty acid) [56]. Such broad reactivity with cannabinoid receptor-insensitive bioactive fatty acid amides should be considered when the molecular mechanisms for the phenotype of FAAH−/− mice are examined. Although FAAH hydrolyzes 2-AG at a high rate [57], its contribution to 2-AG degradation in the brain was estimated to be minor [58, 59]. Considering the physiological importance of FAAH, naturally occurring single-nucleotide polymorphisms were examined in the human FAAH gene. Interestingly, the cytosine 385 to adenine missense mutation was found to be strongly associated with street drug use and problem drug/alcohol use [60]. This mutation results in the substitution of a threonine for Pro129, and the P129T variant showed enhanced sensitivity to proteolytic degradation. Thus, the functional abnormality of FAAH could be linked to drug abuse and dependence in humans.

As FAAH has been suggested to be involved in a variety of symptoms, including anxiety, depression, and neuropathic pain, FAAH inhibitors are expected to be therapeutic drugs [13]. A large number of specific FAAH inhibitors have been developed, and include URB597 [61], OL-135 [62], PF-3845 [63], and PF-04457845 [64]. Furthermore, the full spectrum of cannabimimetic activities was expected with dual inhibition of FAAH and MAGL, the principal enzyme for the degradation of 2-AG. In fact, treatment with fluorophosphonate compounds as the dual inhibitors increased the brain levels of both anandamide and 2-AG more than 10-fold, resulting in the complete appearance of the ‘tetrad of cannabinoid’ (analgesia, hypomotility, hypothermia, and catalepsy) [65]. JZL195, which inhibits both FAAH and MAGL, also mimicked the pharmacological activities of the CB1 receptor agonist in vivo [66].

An isozyme of FAAH with ~ 20% sequence identity at the amino acid level is expressed in humans, but not in rodents [67]. The original FAAH and this newly discovered isozyme were designated FAAH-1 and FAAH-2, respectively. Interestingly, unlike FAAH-1, which was localized in endoplasmic reticulum, FAAH-2 was localized on lipid droplets, and its N-terminal hydrophobic region was identified as a lipid droplet localization sequence, suggesting that the role of FAAH-2 is different from that of FAAH-1 [68]. Recently, a catalytically inactive, truncated variant of FAAH-1 [FAAH-like anandamide transporter (FLAT)] was reported to have the ability to bind to anandamide and drive anandamide transport in neurons [69]. However, this finding has been questioned, for several reasons, including the lack of endogenous FLAT in brain and neural cells, and the remaining FAAH-like catalytic activity of recombinant FLAT [70].

Similarly to the oxygenation of free arachidonic acid to various bioactive eicosanoids, the arachidonic acid chain of anandamide can be oxygenated (Fig. 2) [27, 28]. These metabolites include hydroxyl derivatives of anandamide produced by cytochrome P450 [71], hydroperoxy derivatives produced by lipoxygenases [72, 73], and prostaglandin-like ethanolamides (called prostamides) produced by cyclooxygenase (COX)-2 [74, 75]. Despite their reported biological activities, the physiological significance of these compounds is poorly understood.

Alternative pathways for anandamide and other NAEs

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References

These newly discovered pathways for NAE metabolism include: (a) members of the PLA/acyltransferase (PLA/AT) family; (b) multistep pathways via N-acylated lysophospholipid; and (c) NAAA. These enzymes/pathways were found in animal tissues as possible alternatives to Ca2+-dependent N-acyltransferase, NAPE-PLD, and FAAH, respectively.

PLA/AT family proteins as Ca2+-independent N-acyltransferases

In an attempt to identify Ca2+-dependent N-acyltransferase, we noticed that the reaction catalyzed by N-acyltransferase is similar to that catalyzed by lecithin retinol acyltransferase (LRAT). LRAT transfers an acyl chain from the sn-1 position of lecithin (PtdCho) to retinol (vitamin A alcohol), resulting in the formation of retinyl ester (the storage form of vitamin A). We speculated that the primary structure of N-acyltransferase might be analogous to that of LRAT. LRAT is a member of the NlpC/P60 thiol protease superfamily [76], and has considerable sequence homology with five members (HRASLS1–5) of the HRAS-like suppressor (HRASLS) family (Fig. 5) [77]. HRASLS3 (also referred to as H-rev107) is a representative molecule of this family, and has been analyzed as a class II tumor suppressor that negatively regulates the activity of the oncoprotein Ras [78]. Later, HRASLS1 (A-C1) [79], HRASLS2 [80], and HRASLS4 [tazarotene-induced protein 3 (TIG3)] [81] were also reported to have tumor-suppressing activity. As catalytically important residues, including cysteine and histidine as the putative catalytic dyad, are highly conserved among LRAT and HRASLS family members, HRASLS3 was earlier suggested to be an acyltransferase [76]. However, the enzymatic properties of the family members were not examined until we reported that HRASLS5 (H-rev107-like protein 5) has an N-acyltransferase activity, forming NAPE by transferring an acyl group from PtdCho to PtdEtn [31, 82]. HRASLS5 was present mainly in the cytosolic fraction, and its N-acyltransferase activity was only slightly stimulated by Ca2+. Interestingly, the enzyme removed a fatty acyl group from both the sn-1 and sn-2 positions of the acyl donor PtdCho. These results strongly suggested that HRASLS5 is different from the known membrane-associated Ca2+-dependent N-acyltransferase. Our studies revealed that all other members of the HRASLS family (HRASLS1–4) also have N-acyltransferase activity [77, 83, 84]. Very recently, Golczak et al. also reported that purified HRASLS2, HRASLS3 and HRASLS4 generate NAPE from PtdCho and PtdEtn [85].

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Figure 5. Primary structures of human LRAT and five members of the HRASLS (PLA/AT) family. Closed and shaded boxes indicate identity in all six and any four or five polypeptides, respectively. Dashes denote deletion of amino acids as compared with the other sequences. Asterisks indicate amino acids forming the catalytic triad.

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Importantly, HRASLS1–5 showed not only N-acyltransferase activity, but also PLA1 and PLA2 (PLA1/2) activity, hydrolyzing the sn-1 and sn-2 ester bonds of PtdCho or PtdEtn, and O-acyltransferase activity, transferring an acyl group from PtdCho to the hydroxyl group of lyso-PtdCho. We thus proposed calling HRASLS1–5 PLA/AT-1–5 [77]. PLA/AT-1, PLA/AT-2 and PLA/AT-5 showed relatively high N-acyltransferase activity relative to PLA1/2 activity, whereas PLA/AT-3 and PLA/AT-4 showed lower N-acyltransferase activity [77, 86]. When the homogenate of human embryonic kidney (HEK)293 cells, stably expressing PLA/AT-2, was examined, both the soluble and particulate fractions showed N-acyltransferase activity, with 1.4-fold higher activity in the soluble fraction. The N-acyltransferase activity was not stimulated by Ca2+ in either fraction. Thus, PLA/AT-2 was also presumed to be different from the known Ca2+-dependent N-acyltransferase.

Although PLA/AT proteins show N-acyltransferase activity in vitro, it remained unclear whether these molecules generate NAPE in living cells. When we metabolically labeled COS-7 cells transiently expressing recombinant PLA/AT-1–5 with [14C]ethanolamine, we found that PLA/AT-1, PLA/AT-2, PLA/AT-4 and PLA/AT-5 generated significant amounts of [14C]NAPE and [14C]NAE [86]. Liquid chromatography–tandem MS demonstrated that the stable expression of PLA/AT-2 in HEK293 cells greatly increased endogenous levels of NAPEs and NAEs. Furthermore, additional expression of NAPE-PLD in the PLA/AT-2-expressing cells led to efficient conversion of the increased NAPE to NAE. RT-PCR revealed that human HeLa cells expressed endogenous PLA/AT-2, and the knockdown of this protein by small interfering RNA lowered the endogenous level of NAPE. Taken together, these results suggested that the PLA/AT proteins produce NAPE, which serves as a precursor of NAE in living cells. In the cells overexpressing recombinant PLA/AT proteins, the generation of NAPE proceeded without any cellular stimuli. However, we could not rule out the possibility that the N-acyltransferase activity of native PLA/AT proteins is regulated by intracellular Ca2+ or other signaling molecules.

We reported that PLA/AT-3 (H-rev107) functions mainly as PLA1/2, with low N-acyltransferase activity [83, 84]. However, Duncan et al. termed this molecule adipose-specific PLA2, and classified it as group XVI PLA2 (PLA2G16) [87]. Recently, two groups demonstrated that PLA/AT-3 has PLA1 activity that is as potent as its PLA2 activity [85, 88]. PLA/AT-3 attracted attention because of its abundant expression in adipose tissue, its induction during adipose differentiation, and its ability to suppress lipolysis [89, 90]. We also showed that the overexpression of PLA/AT-3 in HEK293 cells caused dysfunction of peroxisomes and a remarkable decrease in ether-type triglycerides and plasmalogens [91]. It remains to be determined whether the N-acyltransferase activity of PLA/AT proteins is related to their activities regarding tumor suppression, obesity, and dysfunction of peroxisomes. Recently, the N-terminal catalytic domain of PLA/AT-3 was characterized by solution NMR structure analysis [92] and X-ray crystallography [85, 88]. Together with site-directed mutagenesis studies, these studies increased our knowledge of the structure and function of PLA/AT-3, including the following: (a) the catalytic triad in the active site is composed of His23, His35, and Cys113 (His35 is replaced by asparagine in PLA/AT-1); (b) the acylation site in the acyl–protein complex is Cys113; and (c) the C-terminal transmembrane domain is required for the interfacial catalysis.

Multistep pathways via N-acylated lysophospholipid

Whereas NAPE-PLD directly releases NAE from NAPE, multistep pathways for the biosynthesis of NAE from NAPE via N-acylated lysophospholipid were also suggested in the 1980s (Fig. 2) [93]. We showed that the secretory PLA2 isoforms of groups IB, IIA and V hydrolyzed NAPE to 1-acyl-lyso-NAPE, which was further converted to NAE by a membrane-associated lyso-PLD-like enzyme existing in brain and other tissues of rat [94]. Although this ‘lyso-PLD’ was distinguished from NAPE-PLD by its catalytic properties, its further characterization has not been carried out. The presence of NAEs in the tissues of NAPE-PLD−/− mice clarified that NAE can be formed by an alternative pathway(s) in vivo [43]. The double O-deacylation of NAPE via lyso-NAPE and further hydrolysis of the resultant metabolite glycerophospho-NAE (GP-NAE) to NAE and glycerol 3-phosphate were proposed as one alternative pathway (Fig. 2) [95]. GP-NAE was actually detected in mouse brain tissue [96]. In the analysis of NAPE-PLD−/− mice, we found a remarkable increase in endogenous brain levels of lyso-NAPE and GP-NAE as well as of NAPE [44]. These results suggested that NAPE accumulates in the brain because of the deficiency of NAPE-PLD, and is degraded in the alternative pathway via lyso-NAPE and GP-NAE. By the functional proteomic isolation method with fluorophosphonate-biotin probe, α/β-hydrolase 4 [α/β-hydrolase domain-containing protein (ABHD)4] was demonstrated to be responsible for the double O-deacylation that generates GP-NAE from NAPE via lyso-NAPE [95]. ABHD4 thus hydrolyzes both NAPE and lyso-NAPE by PLA1/2 (or PLB) and lysophospholipase activities. ABHD4 preferentially hydrolyzed lyso-NAPE among various lysophospholipids, including lyso-PtdEtn, lyso-PtdCho, and lysophosphatidylserine. Regarding the N-acyl species of lyso-NAPEs, ABHD4 did not distinguish between saturated and polyunsaturated acyl chains. Ser146, in the consensus sequence GXSXG, was presumed to be the catalytic nucleophile. It was recently reported that knockdown of ABHD4 inhibits anoikis (cell death in response to loss of cell–cell and cell–matrix interactions) in prostate epithelial cells [97]. Its relationship to NAE metabolism is unclear.

Glycerophosphodiester phosphodiesterase 1 (GDE1) was shown to hydrolyze GP-NAE to NAE and glycerol 3-phosphate [96]. On the basis of tissue distribution and catalytic properties, including Mg2+ requirement, the brain enzyme activity hydrolyzing GP-NAE to NAE was attributed to GDE1. Thus, the combination of ABHD4 and GDE1 was considered to form a NAPE-PLD-independent pathway. Initially, this pathway was expected to be responsible for anandamide formation, as the brain level of anandamide was not altered in NAPE-PLD−/− mice, in contrast to the decrease in the levels of saturated NAEs [43]. However, neither ABHD4 nor GDE1 showed a preference for N-arachidonoyl species of lyso-NAPE and GP-NAE (precursors of anandamide). It remains unclear whether or not the ABHD4–GDE1 pathway is involved in the selective formation of anandamide. Alternatively, in our analysis of NAPE-PLD−/− mice, the brain level of anandamide was significantly lower, together with the levels of other NAEs [44].

GDE1 is an integral membrane glycoprotein, and was originally discovered as MIR16, a protein interacting with RGS16 (a regulator of G protein signaling) [98]. Later GDE1 was shown to have a phosphodiesterase activity, preferentially hydrolyzing glycerophosphoinositol [99]. GDE1−/− mice were born at the expected Mendelian frequency, were viable and healthy, and showed no abnormal cage behavior [100]. Expectedly, the formation of NAE from GP-NAE or lyso-NAPE was hardly detected in the brain homogenates. However, endogenous brain levels of NAEs were not significantly different between the homozygous and heterozygous mice. Furthermore, no significant difference was seen between NAPE-PLD−/− mice and mice with double knockout of GDE1 and NAPE-PLD. These results suggest that enzymes or pathways other than NAPE-PLD and the ABHD4–GDE1 pathway are involved in NAE formation. Further analysis of GDE1−/− mice demonstrated that brain levels of glycerophosphoinositol, glycerophosphoserine and glycerophosphoglycerate were highly elevated [101]. In agreement with these findings, the brain level of free serine, which should be released from glycerophosphoserine by GDE1, was significantly reduced.

The fact that the brain contains plasmalogen-type ethanolamine phospholipid (plasmenylethanolamine) in abundance [102] suggests that pNAPE also exists in the same tissue and serves as a precursor of NAE (Fig. 6). In rat brain, 65% of N-arachidonoylethanolamine phospholipids were of the plasmalogen type [103]. In agreement with an earlier report using rat heart microsomes [104], recombinant NAPE-PLD hydrolyzed N-palmitoyl-plasmenylethanolamine to PEA at 70% of the rate of N-palmitoyl-PtdEtn hydrolysis [44]. The brain levels of pNAPE and its lyso form (lyso-pNAPE) in NAPE-PLD−/− mice were much higher than those in wild-type mice, as shown by liquid chromatography–tandem MS. Furthermore, the brain homogenate of NAPE-PLD−/− mice converted pNAPE to NAE, and the homogenate also released NAE from lyso-pNAPE. As lyso-pNAPE has a vinyl ether bond rather than an ester bond at the sn-1 position, lysophospholipases such as ABHD4 should be inactive with this lysophospholipid. Therefore, it was likely that a lyso-PLD-type phosphodiesterase directly releases NAE from lyso-pNAPE. The lyso-PLD-type enzyme found in the brain was active at neutral pH, and converted N-palmitoyl-lysoplasmenylethanolamine, N-oleoyl-lysoplasmenylethanolamine and N-arachidonoyl-lysoplasmenylethanolamine to their corresponding NAEs at similar rates. The activity was stimulated by 2 mm Mg2+ and inhibited by 0.1% Triton X-100. We found that recombinant GDE1 showed weak lyso-PLD activity in hydrolyzing N-palmitoyl-lysoplasmenylethanolamine in addition to the aforementioned GP-NAE-hydrolyzing activity. As GDE1 is expressed in brain, GDE1 may be, at least in part, responsible for the brain lyso-PLD activity.

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Figure 6. Biosynthetic pathways of NAE from pNAPE.

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PLC-mediated hydrolysis of NAPE to NAE phosphate and dephosphorylation to generate NAE is another multistep route for the conversion of NAPE to NAE that does not go through lyso-NAPE (Fig. 2). Treatment of RAW264.7 mouse macrophages with LPS potently enhanced anandamide levels, despite the downregulation of NAPE-PLD [39, 105]. Protein tyrosine phosphatase, nonreceptor type 22 was identified as one of the gene products that increase the anandamide level, and exhibited a phosphatase activity that generates anandamide from anandamide phosphate. Src homology 2 domain-containing inositol-5-phosphatase 1 showed the same phosphatase activity. Although the PLC–phosphatase pathway was suggested to function in the brain of NAPE-PLD−/− mice [106], the PLC-like enzyme responsible remains uncharacterized.

NAAA

We found a lysosomal enzyme hydrolyzing NAEs, first in CMK human megakaryoblastic cells [107], and later in the lung and other tissues of rats [108]. cDNA cloning of this enzyme, referred to as NAAA, showed it to be a cysteine hydrolase belonging to the N-terminal nucleophile hydrolase superfamily (Fig. 7) [109-111]. No sequence homology was seen between NAAA and FAAH. Prior to our identification, NAAA was recognized as acid ceramidase-like protein from sequence homology [112]. Acid ceramidase is a lysosomal enzyme that hydrolyzes ceramide to sphingosine and fatty acid. In agreement with its localization in lysosomes, NAAA is active only at acidic pH, and hydrolyzes various NAEs, with a preference for PEA in vitro. Human NAAA is a glycoprotein with four N-glycosylation sites [113, 114]. Like acid ceramidase [115], recombinant NAAA is produced as an inactive proenzyme, and is converted by autocatalytic cleavage between Phe125 and Cys126 to a catalytically active heterodimer composed of α and β subunits. The N-terminal 28 amino acids form a signal peptide, which is not contained in the α subunit [114]. Although the β subunit was purified from rat lung [108], native αβ heterodimer has not been isolated. It remains unclear whether native NAAA stably exists as the heterodimer. On the basis of the sequence homology among the family members, the N-terminal cysteine of the β subunit (Cys126 in human NAAA) has been presumed to be the catalytic nucleophile. This was recently demonstrated by showing that β-lactones, which inhibit NAAA, acylate Cys126 [116, 117].

image

Figure 7. The primary structures of human NAAA and acid ceramidase. Asterisks and dots denote identity and similarity of the two sequences, respectively. Dashes indicate deletion of amino acids as compared with the other sequence. Diamonds and the circle indicate N-glycosylation sites and the catalytic nucleophile of NAAA, respectively. The arrow and the arrowhead denote a site of cleavage by signal peptidase (NAAA) and that between the α and β subunits (both NAAA and acid ceramidase), respectively.

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NAAA is expressed in various human and rodent tissues, with predominant expression in macrophages [118, 119]. The expression of NAAA was highest in the prostate among various human tissues [120]. Nonionic detergents (Triton X-100 and Nonidet P-40) and the thiol-reducing agent dithiothreitol have been used as NAAA stimulators in vitro. As endogenous stimulators, choline-containing or ethanolamine-containing phospholipids (PtdCho, PtdEtn, and sphingomyelin) and dihydrolipoic acid (the reduced form of α-lipoic acid) could substitute for Nonidet P-40 and dithiothreitol, respectively [121]. These endogenous molecules may function by keeping NAAA active in lysosomes. As it is likely that FAAH and NAAA contribute to the degradation of NAEs in vivo, we investigated whether or not compensatory induction of NAAA mRNA occurs in the tissues of FAAH−/− mice. However, the expression levels in various tissues were not significantly different from those in wild-type mice [121].

Considering that NAAA preferentially hydrolyzes PEA over other NAEs, selective NAAA inhibitors that may increase local levels of endogenous PEA were expected to be anti-inflammatory and analgesic drugs [122]. To date, many compounds have been reported to selectively inhibit NAAA (Fig. 8) [117, 123-130]. The most potent NAAA inhibitors thus far reported are lactone derivatives such as (S)-N-(2-oxo-3-oxetanyl)-3-phenylpropionamide [(S)-OOPP], (S)-N-(2-oxo-3-oxetanyl)biphenyl-4-carboxamide, and (2S,3R)-2-methyl-4-oxo-3-oxetanylcarbamic acid 5-phenylpentyl ester (URB913/ARN077), whose IC50 values were 420, 115 and 127 nm, respectively [125, 126, 128]. (S)-OOPP normalized the decreased PEA levels in carrageenan-stimulated leukocytes and LPS-treated RAW264.7 macrophage cells, and led to a reduction in neutrophil migration and inhibition of carrageenan-induced plasma extravasation [125]. Recently, we reported that lipophilic amines such as pentadecylamine and tridecyl 2-aminoacetate inhibited NAAA with IC50 values of 5.7 and 11.8 μm, respectively, and showed much weaker effects on FAAH [129]. These simple structures may provide a scaffold for further improvement.

image

Figure 8. Chemical structures of specific NAAA inhibitors. (A) Lactone derivatives. (B) Other compounds. IC50 values are shown in parentheses.

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Metabolism of 2-AG

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References

Biosynthesis of 2-AG

Earlier, the PLC–diacylglycerol lipase (DAGL) pathway was considered to play a role in the release of arachidonic acid from phospholipid [131]. This pathway is now known to be the most important route in the biosynthesis of 2-AG (Fig. 3) [11, 132]. It is generally accepted that 2-AG is formed through this pathway in postsynaptic neurons in response to depolarization and stimulation of Gq/11-coupled receptors. The resultant 2-AG is released, and mediates retrograde synaptic suppression through CB1 receptors at presynaptic terminals [133]. In this pathway, PLC hydrolyzes 2-arachidonoyl-phosphatidylinositol (PtdIns) to arachidonic acid-containing diacylglycerol (DAG), which is subsequently hydrolyzed to 2-AG by DAGL. The most abundant PtdIns molecule in mammalian tissues is 1-stearoyl-2-arachidonoyl-PtdIns, enabling the effective generation of 2-AG over other MAGs. Among the β, γ, δ, ε, ζ, and η subtypes of PLC, the β subtype is characterized by the stimulation by Gq/11-coupled receptors [134], which suggests that it has a central role in neurotransmitter-dependent 2-AG formation mediated by Gq/11-coupled receptors in postsynaptic neurons. In fact, each isoform (β1–β4) of PLCβ, which is expressed regiospecifically in different brain areas, is stimulated by different Gq/11-coupled receptors, including the group I metabotropic glutamate receptors (mGluRs) mGluR1 and mGluR5, and the muscarinic acetylcholine receptor M1 [133]. PA [135] and PtdCho [136] may also be converted to 2-arachidonoyl-DAG, which, in turn, serves as a precursor of 2-AG. Another pathway initiated from PtdIns leads to the formation of lyso-PtdIns by PLA1, followed by the release of 2-AG by lyso-PtdIns-specific PLC (Fig. 3) [137, 138]. Furthermore, 2-AG can be formed from 2-arachidonoyl-lysophosphatidic acid (LPA) (Fig. 3) [139].

DAGL, a membrane-associated enzyme hydrolyzing DAG preferentially at the sn-1 position, was first reported in 1981 [140, 141]. cDNA cloning revealed that human DAGL has two closely related genes, α and β, a lipase-3 motif, a serine lipase motif, and four putative transmembrane domains [142]. DAGLα was widely distributed in human and mouse tissues, with high expression levels in the nervous system (human and mouse) and pancreas (human). DAGLα was colocalized with Gq/11-coupled receptors, Gq/11α and PLCβ at particular synaptic and neuronal surfaces [133]. The knockdown of DAGLα by RNA interference suggested its involvement in the mGluR-dependent formation of 2-AG in neuroblastoma cells [143]. Both DAGLα-deficient and DAGLβ-deficient mice were viable, and their general appearance was normal [144, 145]. DAGLα-deficient mice showed a remarkable reduction in brain 2-AG levels, and failed in Ca2+-dependent and Gq/11-coupled receptor-driven retrograde synaptic suppression. These results strongly suggested the involvement of DAGLα in the generation of 2-AG, which is responsible for CB1 receptor-dependent retrograde signaling. In contrast, the brain 2-AG levels of DAGLβ-deficient mice were normal [144] or reduced by 50% [145]. The classical DAGL inhibitors, RHC80267 and tetrahydrolipstatin, inhibited both DAGL isoforms, and lowered endogenous 2-AG levels [146]. More selective DAGL inhibitors include O-3841, OMDM-188, and O-5596 [147-149].

Degradation of 2-AG

The major degradative pathway of 2-AG is its hydrolysis to arachidonic acid and glycerol [11, 150] (Fig. 3). The hydrolysis of 2-AG in brain is mainly catalyzed by MAGL, and < 15% of the activity is attributed to other hydrolases, such as ABHD6, ABHD12, and FAAH [59]. Mammalian MAGL was purified in 1976 [151] and cloned in 1997 [152]. The enzyme hydrolyzes not only 2-AG but also other 2-MAGs and 1-MAGs, and it is a soluble protein that associates with membranes in a peripheral manner [153]. A serine hydrolase belonging to the α/β-hydrolase superfamily, MAGL has a catalytic triad composed of Ser122, Asp239, and His269. MAGL crystallized as a dimer, and its three-dimensional structure has been analyzed in detail [154, 155]. MAGL is expressed in many tissues, and plays an important role in the degradation of 2-AG that is responsible for endocannabinoid signaling in the brain [152, 156]. In the cerebellum, the expression of MAGL was high within parallel fiber terminals, weak in Bergman glia, and absent in other synaptic terminals. However, 2-AG was broadly degraded in a synapse-type-independent manner by MAGL [157].

MAGL inhibitors have attracted much attention as promising therapeutic drugs and pharmacological probes, owing to their ability to enhance 2-AG signaling [13, 150, 153, 158]. A large number of sulfhydryl blockers and serine hydrolase inhibitors have been reported to be nonspecific or specific MAGL inhibitors. An O-aryl carbamate, JZL184, is one of the most potent inhibitors selective for MAGL. In JZL184-treated mice, the brain level of 2-AG was increased, and CB1-dependent analgesia, hypothermia and hypomotility appeared [159]. KML29, a recently developed derivative of JZL184, was more selective for MAGL, and did not show detectable crossreactivity with FAAH [160]. In MAGL-deficient mice, the brain 2-AG-hydrolyzing activity dramatically decreased and the brain 2-AG level was highly elevated [161-163]. Notably, similarly to mice chronically treated with JZL184, MAGL-deficient mice showed tolerance to CB1 agonists. Downregulation and desensitization of brain CB1 were also observed. Recent studies have revealed roles for MAGL other than 2-AG degradation. For example, MAGL deficiency impaired lipolysis and attenuated diet-induced insulin resistance [163]. Arachidonic acid formation as a result of MAGL generates prostaglandins that promote neuroinflammation in the brain [164]. Moreover, MAGL was highly expressed in aggressive human cancer cells and primary tumors. Its overexpression in nonaggressive cancer cells increased their pathogenicity phenotypes [165].

ABHD6 and ABHD12 are additional MAG hydrolases belonging to the α/β-hydrolase superfamily, with the postulated catalytic triad serine–aspartic acid–histidine [166, 167]. ABHD6 was notable for its expression in the mouse microglial cell line BV-2, in which MAGL was not expressed [168]. In adult mouse cortex, ABHD6 was located postsynaptically in neurons, and it was suggested to constitute a rate-limiting step in 2-AG signaling. UCM710 inhibited FAAH and ABHD6, but not MAGL [169]. Mutations in the ABHD12 gene are causally linked to a neurodegenerative disease termed polyneuropathy, hearing loss, ataxia, retinitis pigmentosa, and cataract (PHARC), suggesting essential functions of ABHD12 in the nervous system and eye [170].

As with anandamide, the arachidonoyl moiety of 2-AG can be oxygenated by COX-2 and lipoxygenases, resulting in the formation of glyceryl prostaglandins or hydroperoxy derivatives of 2-AG, respectively (Fig. 3) [171, 172]. Some biological activities of glyceryl prostaglandins have been reported [172]. Interestingly, R-enantiomers of ibuprofen, naproxen and flurbiprofen selectively inhibited the endocannabinoid oxygenation catalyzed by COX-2, but were inactive with free arachidonic acid as a COX-2 substrate. Thus, these inhibitors were considered to be substrate-selective inhibitors of endocannabinoid oxygenation [173].

Conclusions

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References

We have discussed new enzymes and pathways that may be involved in the metabolism of endocannabinoids and related NAEs. The new enzymes/pathways in the metabolism of anandamide and other bioactive NAEs included: (a) the members of the HRASLS family or PLA/AT family, which function as Ca2+-independent N-acyltransferases; (b) NAPE-PLD-independent multistep pathways via N-acylated lysophospholipid; and (c) NAAA, a lysosomal enzyme that preferentially hydrolyzes PEA. Although their physiological significance is poorly understood, these new enzymes/pathways may serve as novel targets for the development of therapeutic drugs. For example, selective NAAA inhibitors may be promising as novel anti-inflammatory and analgesic drugs. Ca2+-dependent N-acyltransferase in the canonical pathway remains molecularly uncharacterized. The identification of the gene is eagerly anticipated. Recent studies enabled the analysis of genetic and pharmacological deficiency of 2-AG-metabolizing enzymes, and confirmed the essential role of 2-AG in the mediation of retrograde synaptic suppression. Elucidation of the physiological functions of ABHD6 and ABHD12 is also expected. After the submission of this manuscript, Blankman et al. reported that ABHD12 is a principal lysophosphatidylserine lipase in the mammalian brain [174]. The mutations in ABHD12 caused the human neurodegenerative disorder PHARC, and the phenotypes of ABHD12−/− mice resembled those of human PHARC patients.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References

We are grateful to D. G. Deutsch (Department of Biochemistry and Cell Biology, Stony Brook University, Stony Brook, NY, USA) for critical reading of this manuscript.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Canonical pathways for anandamide and other NAEs
  5. Alternative pathways for anandamide and other NAEs
  6. Metabolism of 2-AG
  7. Conclusions
  8. Acknowledgements
  9. References
  • 1
    Pacher P, Bátkai S & Kunos G (2006) The endocannabinoid system as an emerging target of pharmacotherapy. Pharmacol Rev 58, 389462.
  • 2
    Pertwee RG, Howlett AC, Abood ME, Alexander SP, Di Marzo V, Elphick MR, Greasley PJ, Hansen HS, Kunos G, Mackie K, et al. (2010) International union of basic and clinical pharmacology. LXXIX. Cannabinoid receptors and their ligands: beyond CB1 and CB2. Pharmacol Rev 62, 588631.
  • 3
    Devane WA, Hanus L, Breuer A, Pertwee RG, Stevenson LA, Griffin G, Gibson D, Mandelbaum A, Etinger A & Mechoulam R (1992) Isolation and structure of a brain constituent that binds to the cannabinoid receptor. Science 258, 19461949.
  • 4
    Mechoulam R, Ben-Shabat S, Hanus L, Ligumsky M, Kaminski NE, Schatz AR, Gopher A, Almog S, Martin BR, Compton DR, et al. (1995) Identification of an endogenous 2-monoglyceride, present in canine gut, that binds to cannabinoid receptors. Biochem Pharmacol 50, 8390.
  • 5
    Sugiura T, Kondo S, Sukagawa A, Nakane S, Shinoda A, Itoh K, Yamashita A & Waku K (1995) 2-Arachidonoylglycerol: a possible endogenous cannabinoid receptor ligand in brain. Biochem Biophys Res Commun 215, 8997.
  • 6
    Hansen HS & Diep TA (2009) N-Acylethanolamines, anandamide and food intake. Biochem Pharmacol 78, 553560.
  • 7
    LoVerme J, La Rana G, Russo R, Calignano A & Piomelli D (2005) The search for the palmitoylethanolamide receptor. Life Sci 77, 16851698.
  • 8
    Pavón FJ, Serrano A, Romero-Cuevas M, Alonso M & Rodríguez de Fonseca F (2010) Oleoylethanolamide: a new player in peripheral control of energy metabolism. Therapeutic implications. Drug Discov Today Dis Mech 7, e175e183.
  • 9
    Hansen KB, Rosenkilde MM, Knop FK, Wellner N, Diep TA, Rehfeld JF, Andersen UB, Holst JJ & Hansen HS (2011) 2-Oleoyl glycerol is a GPR119 agonist and signals GLP-1 release in humans. J Clin Endocrinol Metab 96, E14091417.
  • 10
    Ueda N, Tsuboi K & Uyama T (2010a) Enzymological studies on the biosynthesis of N-acylethanolamines. Biochim Biophys Acta 1801, 12741285.
  • 11
    Ueda N, Tsuboi K, Uyama T & Ohnishi T (2011) Biosynthesis and degradation of the endocannabinoid 2-arachidonoylglycerol. BioFactors 37, 17.
  • 12
    Ueda N & Tsuboi K (2012) Discrimination between two endocannabinoids. Chem Biol 19, 545547.
  • 13
    Petrosino S & Di Marzo V (2010) FAAH and MAGL inhibitors: therapeutic opportunities from regulating endocannabinoid levels. Curr Opin Investig Drugs 11, 5162.
  • 14
    Elphick MR & Egertová M (2001) The neurobiology and evolution of cannabinoid signalling. Philos Trans R Soc Lond B Biol Sci 356, 381408.
  • 15
    McPartland JM, Matias I, Di Marzo V & Glass M (2006) Evolutionary origins of the endocannabinoid system. Gene 370, 6474.
  • 16
    Kilaru A, Blancaflor EB, Venables BJ, Tripathy S, Mysore KS & Chapman KD (2007) The N-acylethanolamine-mediated regulatory pathway in plants. Chem Biodivers 4, 19331955.
  • 17
    Lucanic M, Held JM, Vantipalli MC, Klang IM, Graham JB, Gibson BW, Lithgow GJ & Gill MS (2011) N-acylethanolamine signalling mediates the effect of diet on lifespan in Caenorhabditis elegans. Nature 473, 226229.
  • 18
    Coulon D, Faure L, Salmon M, Wattelet V & Bessoule JJ (2012a) N-Acylethanolamines and related compounds: aspects of metabolism and functions. Plant Sci 184, 129140.
  • 19
    Schmid HHO, Schmid PC & Natarajan V (1990) N-Acylated glycerophospholipids and their derivatives. Prog Lipid Res 29, 143.
  • 20
    Schmid HHO (2000) Pathways and mechanisms of N-acylethanolamine biosynthesis: can anandamide be generated selectively? Chem Phys Lipids 108, 7187.
  • 21
    Wellner N, Diep TA, Janfelt C & Hansen HS (2013) N-Acylation of phosphatidylethanolamine and its biological functions in mammals. Biochim Biophys Acta 1831, 652662.
  • 22
    Coulon D, Faure L, Salmon M, Wattelet V & Bessoule JJ (2012b) Occurrence, biosynthesis and functions of N-acylphosphatidylethanolamines (NAPE): not just precursors of N-acylethanolamines (NAE). Biochimie 94, 7585.
  • 23
    Gillum MP, Zhang D, Zhang X-M, Erion DM, Jamison RA, Choi C, Dong J, Shanabrough M, Duenas HR, Frederick DW, et al. (2008) N-Acylphosphatidylethanolamine, a gut-derived circulating factor induced by fat ingestion, inhibits food intake. Cell 135, 813824.
  • 24
    Wellner N, Tsuboi K, Madsen AN, Holst B, Diep TA, Nakao M, Tokumura A, Burns MP, Deutsch DG, Ueda N, et al. (2011) Studies on the anorectic effect of N-acylphosphatidylethanolamine and phosphatidylethanolamine in mice. Biochim Biophys Acta 1811, 508512.
  • 25
    Deutsch DG, Ueda N & Yamamoto S (2002) The fatty acid amide hydrolase (FAAH). Prostaglandins Leukot Essent Fatty Acids 66, 201210 (erratum appears in Prostaglandins Leukot Essent Fatty Acids 68, 69).
  • 26
    McKinney MK & Cravatt BF (2005) Structure and function of fatty acid amide hydrolase. Annu Rev Biochem 74, 411432.
  • 27
    Vandevoorde S & Lambert DM (2007) The multiple pathways of endocannabinoid metabolism: a zoom out. Chem Biodivers 4, 18581881.
  • 28
    Fowler CJ (2007) The contribution of cyclooxygenase-2 to endocannabinoid metabolism and action. Br J Pharmacol 152, 594601.
  • 29
    Hansen HS, Moesgaard B, Hansen HH & Petersen G (2000) N-Acylethanolamines and precursor phospholipids – relation to cell injury. Chem Phys Lipids 108, 135150.
  • 30
    Cadas H, di Tomaso E & Piomelli D (1997) Occurrence and biosynthesis of endogenous cannabinoid precursor, N-arachidonoyl phosphatidylethanolamine, in rat brain. J Neurosci 17, 12261242.
  • 31
    Jin X-H, 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. J Biol Chem 282, 36143623.
  • 32
    Schmid HHO, Schmid PC & Berdyshev EV (2002) Cell signaling by endocannabinoids and their congeners: questions of selectivity and other challenges. Chem Phys Lipids 121, 111134.
  • 33
    Janfelt C, Wellner N, Leger PL, Kokesch-Himmelreich J, Hansen SH, Charriaut-Marlangue C & Hansen HS (2012) Visualization by mass spectrometry of 2-dimensional changes in rat brain lipids, including N-acylphosphatidylethanolamines, during neonatal brain ischemia. FASEB J 26, 26672673.
  • 34
    Okamoto Y, Wang J, Morishita J & Ueda N (2007) Biosynthetic pathways of the endocannabinoid anandamide. Chem Biodivers 4, 18421857.
  • 35
    Okamoto Y, Morishita J, Tsuboi K, Tonai T & Ueda N (2004) Molecular characterization of a phospholipase D generating anandamide and its congeners. J Biol Chem 279, 52985305.
  • 36
    Wang J, Okamoto Y, Morishita J, Tsuboi K, Miyatake A & Ueda N (2006) Functional analysis of the purified anandamide-generating phospholipase D as a member of the metallo-β-lactamase family. J Biol Chem 281, 1232512335.
  • 37
    Ueda N, Liu Q & Yamanaka K (2001a) Marked activation of the N-acylphosphatidylethanolamine-hydrolyzing phosphodiesterase by divalent cations. Biochim Biophys Acta 1532, 121127.
  • 38
    Wang J, Okamoto Y, Tsuboi K & Ueda N (2008a) The stimulatory effect of phosphatidylethanolamine on N-acylphosphatidylethanolamine-hydrolyzing phospholipase D (NAPE-PLD). Neuropharmacology 54, 815.
  • 39
    Liu J, Wang L, Harvey-White J, Osei-Hyiaman D, Razdan R, Gong Q, Chan AC, Zhou Z, Huang BX, Kim HY, et al. (2006) A biosynthetic pathway for anandamide. Proc Natl Acad Sci USA 103, 1334513350.
  • 40
    Zhu C, Solorzano C, Sahar S, Realini N, Fung E, Sassone-Corsi P & Piomelli D (2011) Proinflammatory stimuli control N-acylphosphatidylethanolamine-specific phospholipase D expression in macrophages. Mol Pharmacol 79, 786792.
  • 41
    Morishita J, Okamoto Y, Tsuboi K, Ueno M, Sakamoto H, Maekawa N & Ueda N (2005) Regional distribution and age-dependent expression of N-acylphosphatidylethanolamine-hydrolyzing phospholipase D in rat brain. J Neurochem 94, 753762.
  • 42
    Moesgaard B, Petersen G, Jaroszewski JW & Hansen HS (2000) Age dependent accumulation of N-acyl-ethanolamine phospholipids in ischemic rat brain: a 31P NMR and enzyme activity study. J Lipid Res 41, 985990.
  • 43
    Leung D, Saghatelian A, Simon GM & Cravatt BF (2006) Inactivation of N-acyl phosphatidylethanolamine phospholipase D reveals multiple mechanisms for the biosynthesis of endocannabinoids. Biochemistry 45, 47204726.
  • 44
    Tsuboi K, Okamoto Y, Ikematsu N, Inoue M, Shimizu Y, Uyama T, Wang J, Deutsch DG, Burns MP, Ulloa NM, et al. (2011) Enzymatic formation of N-acylethanolamines from N-acylethanolamine plasmalogen through N-acylphosphatidylethanolamine-hydrolyzing phospholipase D-dependent and -independent pathways. Biochim Biophys Acta 1811, 565577.
  • 45
    Wangensteen T, Akselsen H, Holmen J, Undlien D & Retterstøl L (2011) A common haplotype in NAPEPLD is associated with severe obesity in a Norwegian population-based cohort (the HUNT study). Obesity 19, 612617.
  • 46
    Sugiura T, Kondo S, Sukagawa A, Tonegawa T, Nakane S, Yamashita A, Ishima Y & 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, 5362.
  • 47
    Arreaza G, Devane WA, Omeir RL, Sajnani G, Kunz J, Cravatt BF & Deutsch DG (1997) The cloned rat hydrolytic enzyme responsible for the breakdown of anandamide also catalyzes its formation via the condensation of arachidonic acid and ethanolamine. Neurosci Lett 234, 5962.
  • 48
    Kurahashi Y, Ueda N, Suzuki H, Suzuki M & Yamamoto S (1997) Reversible hydrolysis and synthesis of anandamide demonstrated by recombinant rat fatty-acid amide hydrolase. Biochem Biophys Res Commun 237, 512515.
  • 49
    Katayama K, Ueda N, Katoh I & Yamamoto S (1999) Equilibrium in the hydrolysis and synthesis of cannabimimetic anandamide demonstrated by a purified enzyme. Biochim Biophys Acta 1440, 205214.
  • 50
    Mukhopadhyay B, Cinar R, Yin S, Liu J, Tam J, Godlewski G, Harvey-White J, Mordi I, Cravatt BF, Lotersztajn S, et al. (2011) Hyperactivation of anandamide synthesis and regulation of cell-cycle progression via cannabinoid type 1 (CB1) receptors in the regenerating liver. Proc Natl Acad Sci USA 108, 63236328.
  • 51
    McCue JM, Driscoll WJ & Mueller GP (2009) In vitro synthesis of arachidonoyl amino acids by cytochrome c. Prostaglandins Other Lipid Mediat 90, 4248.
  • 52
    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, 8387.
  • 53
    McKinney MK & Cravatt BF (2003) Evidence for distinct roles in catalysis for residues of the serine-serine-lysine catalytic triad of fatty acid amide hydrolase. J Biol Chem 278, 3739337399.
  • 54
    Cravatt BF, Demarest K, Patricelli M, Bracey MH, Giang DK, Martin BR & Lichtman AH (2001) Supersensitivity to anandamide and enhanced endogenous cannabinoid signaling in mice lacking fatty acid amide hydrolase. Proc Natl Acad Sci USA 98, 93719376.
  • 55
    Clement AB, Hawkins EG, Lichtman AH & Cravatt BF (2003) Increased seizure susceptibility and proconvulsant activity of anandamide in mice lacking fatty acid amide hydrolase. J Neurosci 23, 39163923.
  • 56
    Saghatelian A, Trauger SA, Want EJ, Hawkins EG, Siuzdak G & Cravatt BF (2004) Assignment of endogenous substrates to enzymes by global metabolite profiling. Biochemistry 43, 1433214339.
  • 57
    Goparaju SK, Ueda N, Yamaguchi H & Yamamoto S (1998) Anandamide amidohydrolase reacting with 2-arachidonoylglycerol, another cannabinoid receptor ligand. FEBS Lett 422, 6973.
  • 58
    Goparaju SK, Ueda N, Taniguchi K & Yamamoto S (1999) Enzymes of porcine brain hydrolyzing 2-arachidonoylglycerol, an endogenous ligand of cannabinoid receptors. Biochem Pharmacol 57, 417423.
  • 59
    Blankman JL, Simon GM & Cravatt BF (2007) A comprehensive profile of brain enzymes that hydrolyze the endocannabinoid 2-arachidonoylglycerol. Chem Biol 14, 13471356.
  • 60
    Sipe JC, Chiang K, Gerber AL, Beutler E & Cravatt BF (2002) A missense mutation in human fatty acid amide hydrolase associated with problem drug use. Proc Natl Acad Sci USA 99, 83948399.
  • 61
    Kathuria S, Gaetani S, Fegley D, Valiño F, Duranti A, Tontini A, Mor M, Tarzia G, La Rana G, Calignano A, et al. (2003) Modulation of anxiety through blockade of anandamide hydrolysis. Nat Med 9, 7681.
  • 62
    Lichtman AH, Leung D, Shelton CC, Saghatelian A, Hardouin C, Boger DL & Cravatt BF (2004) Reversible inhibitors of fatty acid amide hydrolase that promote analgesia: evidence for an unprecedented combination of potency and selectivity. J Pharmacol Exp Ther 311, 441448.
  • 63
    Ahn K, Johnson DS, Mileni M, Beidler D, Long JZ, McKinney MK, Weerapana E, Sadagopan N, Liimatta M, Smith SE, et al. (2009) Discovery and characterization of a highly selective FAAH inhibitor that reduces inflammatory pain. Chem Biol 16, 411420.
  • 64
    Ahn K, Smith SE, Liimatta MB, Beidler D, Sadagopan N, Dudley DT, Young T, Wren P, Zhang Y, Swaney S, et al. (2011) Mechanistic and pharmacological characterization of PF-04457845: a highly potent and selective fatty acid amide hydrolase inhibitor that reduces inflammatory and noninflammatory pain. J Pharmacol Exp Ther 338, 114124.
  • 65
    Nomura DK, Blankman JL, Simon GM, Fujioka K, Issa RS, Ward AM, Cravatt BF & Casida JE (2008) Activation of the endocannabinoid system by organophosphorus nerve agents. Nat Chem Biol 4, 373378.
  • 66
    Long JZ, Nomura DK, Vann RE, Walentiny DM, Booker L, Jin X, Burston JJ, Sim-Selley LJ, Lichtman AH, Wiley JL, et al. (2009a) Dual blockade of FAAH and MAGL identifies behavioral processes regulated by endocannabinoid crosstalk in vivo. Proc Natl Acad Sci USA 106, 2027020275.
  • 67
    Wei BQ, Mikkelsen TS, McKinney MK, Lander ES & Cravatt BF (2006) A second fatty acid amide hydrolase with variable distribution among placental mammals. J Biol Chem 281, 3656936578.
  • 68
    Kaczocha M, Glaser ST, Chae J, Brown DA & Deutsch DG (2010) Lipid droplets are novel sites of N-acylethanolamine inactivation by fatty acid amide hydrolase-2. J Biol Chem 285, 27962806.
  • 69
    Fu J, Bottegoni G, Sasso O, Bertorelli R, Rocchia W, Masetti M, Guijarro A, Lodola A, Armirotti A, Garau G, et al. (2011) A catalytically silent FAAH-1 variant drives anandamide transport in neurons. Nat Neurosci 15, 6469.
  • 70
    Leung K, Vivieca S, Sun J, Glaser ST, Deutsch DG & Kaczocha M (2012) FLAT is not an intracellular anandamide transporter. 22nd Annual Symposium on the Cannabinoids, International Cannabinoid Research Society, Research Triangle Park, NC, USA, pp. 59.
  • 71
    Bornheim LM, Kim KY, Chen B & Correia MA (1993) The effect of cannabidiol on mouse hepatic microsomal cytochrome P450-dependent anandamide metabolism. Biochem Biophys Res Commun 197, 740746.
  • 72
    Ueda N, Yamamoto K, Yamamoto S, Tokunaga T, Shirakawa E, Shinkai H, Ogawa M, Sato T, Kudo I, Inoue K, et al. (1995) Lipoxygenase-catalyzed oxygenation of arachidonylethanolamide, a cannabinoid receptor agonist. Biochim Biophys Acta 1254, 127134.
  • 73
    Hampson AJ, Hill WAG, Zan-Phillips M, Makriyannis A, Leung E, Eglen RM & Bornheim LM (1995) Anandamide hydroxylation by brain lipoxygenase: metabolite structures and potencies at the cannabinoid receptor. Biochim Biophys Acta 1259, 173179.
  • 74
    Yu M, Ives D & Ramesha CS (1997) Synthesis of prostaglandin E2 ethanolamide from anandamide by cyclooxygenase-2. J Biol Chem 272, 2118121186.
  • 75
    Kozak KR, Crews BC, Morrow JD, Wang LH, Ma YH, Weinander R, Jakobsson PJ & Marnett LJ (2002) Metabolism of the endocannabinoids, 2-arachidonylglycerol and anandamide, into prostaglandin, thromboxane, and prostacyclin glycerol esters and ethanolamides. J Biol Chem 277, 4487744885.
  • 76
    Anantharaman V & Aravind L (2003) Evolutionary history, structural features and biochemical diversity of the NlpC/P60 superfamily of enzymes. Genome Biol 4, R11.
  • 77
    Shinohara N, Uyama T, Jin X-H, Tsuboi K, Tonai T, Houchi H & Ueda N (2011) Enzymological analysis of the tumor suppressor A-C1 reveals a novel group of phospholipid-metabolizing enzymes. J Lipid Res 52, 19271935.
  • 78
    Hajnal A, Klemenz R & Schäfer R (1994) Subtraction cloning of H-rev107, a gene specifically expressed in H-ras resistant fibroblasts. Oncogene 9, 479490.
  • 79
    Akiyama H, Hiraki Y, Noda M, Shigeno C, Ito H & Nakamura T (1999) Molecular cloning and biological activity of a novel Ha-Ras suppressor gene predominantly expressed in skeletal muscle, heart, brain, and bone marrow by differential display using clonal mouse EC cells, ATDC5. J Biol Chem 274, 3219232197.
  • 80
    Shyu R-Y, Hsieh Y-C, Tsai F-M, Wu C-C & Jiang S-Y (2009) Cloning and functional characterization of the HRASLS2 gene. Amino Acids 35, 129137.
  • 81
    Disepio D, Ghosn C, Eckert RL, Deucher A, Robinson N, Duvic M, Chandraratna RA & Nagpal S (1998) Identification and characterization of a retinoid-induced class II tumor suppressor/growth regulatory gene. Proc Natl Acad Sci USA 95, 1481114815.
  • 82
    Jin X-H, Uyama T, Wang J, Okamoto Y, Tonai T & Ueda N (2009) cDNA cloning and characterization of human and mouse Ca2+-independent phosphatidylethanolamine N-acyltransferases. Biochim Biophys Acta 1791, 3238.
  • 83
    Uyama T, Morishita J, Jin X-H, Okamoto Y, Tsuboi K & Ueda N (2009a) The tumor suppressor gene H-Rev107 functions as a novel Ca2+-independent cytosolic phospholipase A1/2 of the thiol hydrolase-type. J Lipid Res 50, 685693.
  • 84
    Uyama T, Jin X-H, Tsuboi K, Tonai T & Ueda N (2009b) Characterization of the human tumor suppressors TIG3 and HRASLS2 as phospholipid-metabolizing enzymes. Biochim Biophys Acta 1791, 11141124.
  • 85
    Golczak M, Kiser PD, Sears AE, Lodowski DT, Blaner WS & Palczewski K (2012) Structural basis for the acyltransferase activity of lecithin:retinol acyltransferase-like proteins. J Biol Chem 287, 2379023807.
  • 86
    Uyama T, Ikematsu N, Inoue M, Shinohara N, Jin X-H, Tsuboi K, Tonai T, Tokumura A & Ueda N (2012a) Generation of N-acylphosphatidylethanolamine by members of the phospholipase A/acyltransferase (PLA/AT) family. J Biol Chem 287, 3190531919.
  • 87
    Duncan RE, Sarkadi-Nagy E, Jaworski K, Ahmadian M & Sul HS (2008) Identification and functional characterization of adipose-specific phospholipase A2 (AdPLA). J Biol Chem 283, 2542825436.
  • 88
    Pang XY, Cao J, Addington L, Lovell S, Battaile KP, Zhang N, Rao JL, Dennis EA & Moise AR (2012) Structure/function relationships of adipose phospholipase A2 containing a Cys-His-His catalytic triad. J Biol Chem 287, 3526035274.
  • 89
    Hummasti S, Hong C, Bensinger SJ & Tontonoz P (2008) HRASLS3 is a PPARγ-selective target gene that promotes adipocyte differentiation. J Lipid Res 49, 25352544.
  • 90
    Jaworski K, Ahmadian M, Duncan RE, Sarkadi-Nagy E, Varady KA, Hellerstein MK, Lee HY, Samuel VT, Shulman GI, Kim KH, et al. (2009) AdPLA ablation increases lipolysis and prevents obesity induced by high-fat feeding or leptin deficiency. Nat Med 15, 159168.
  • 91
    Uyama T, Ichi I, Kono N, Inoue A, Tsuboi K, Jin X-H, Araki N, Aoki J, Arai H & Ueda N (2012b) Regulation of peroxisomal lipid metabolism by catalytic activity of tumor suppressor H-rev107. J Biol Chem 287, 27062718.
  • 92
    Ren X, Lin J, Jin C & Xia B (2010) Solution structure of the N-terminal catalytic domain of human H-REV107 – a novel circular permutated NlpC/P60 domain. FEBS Lett 584, 42224226.
  • 93
    Natarajan V, Schmid PC, Reddy PV & Schmid HHO (1984) Catabolism of N-acylethanolamine phospholipids by dog brain preparations. J Neurochem 42, 16131619.
  • 94
    Sun Y-X, Tsuboi K, Okamoto Y, Tonai T, Murakami M, Kudo I & Ueda N (2004) Biosynthesis of anandamide and N-palmitoylethanolamine by sequential actions of phospholipase A2 and lysophospholipase D. Biochem J 380, 749756.
  • 95
    Simon GM & Cravatt BF (2006) Endocannabinoid biosynthesis proceeding through glycerophospho-N-acyl ethanolamine and a role for α/β-hydrolase 4 in this pathway. J Biol Chem 281, 2646526472.
  • 96
    Simon GM & Cravatt BF (2008) Anandamide biosynthesis catalyzed by the phosphodiesterase GDE1 and detection of glycerophospho-N-acyl ethanolamine precursors in mouse brain. J Biol Chem 283, 93419349.
  • 97
    Simpson CD, Hurren R, Kasimer D, MacLean N, Eberhard Y, Ketela T, Moffat J & Schimmer AD (2012) A genome wide shRNA screen identifies α/β hydrolase domain containing 4 (ABHD4) as a novel regulator of anoikis resistance. Apoptosis 17, 666678.
  • 98
    Zheng B, Chen D & Farquhar MG (2000) MIR16, a putative membrane glycerophosphodiester phosphodiesterase, interacts with RGS16. Proc Natl Acad Sci USA 97, 39994004.
  • 99
    Zheng B, Berrie CP, Corda D & Farquhar MG (2003) GDE1/MIR16 is a glycerophosphoinositol phosphodiesterase regulated by stimulation of G protein-coupled receptors. Proc Natl Acad Sci USA 100, 17451750.
  • 100
    Simon GM & Cravatt BF (2010) Characterization of mice lacking candidate N-acyl ethanolamine biosynthetic enzymes provides evidence for multiple pathways that contribute to endocannabinoid production in vivo. Mol BioSyst 6, 14111418.
  • 101
    Kopp F, Komatsu T, Nomura DK, Trauger SA, Thomas JR, Siuzdak G, Simon GM & Cravatt BF (2010) The glycerophospho metabolome and its influence on amino acid homeostasis revealed by brain metabolomics of GDE1(–/–) mice. Chem Biol 17, 831840.
  • 102
    Tsuboi K, Ikematsu N, Uyama T, Deutsch DG, Tokumura A & Ueda N (2013) Biosynthetic pathways of bioactive N-acylethanolamines in brain. CNS Neurol Disord Drug Targets, Epub-20130204-9.
  • 103
    Astarita G, Ahmed F & Piomelli D (2008) Identification of biosynthetic precursors for the endocannabinoid anandamide in the rat brain. J Lipid Res 49, 4857.
  • 104
    Schmid PC, Reddy PV, Natarajan V & Schmid HHO (1983) Metabolism of N-acylethanolamine phospholipids by a mammalian phosphodiesterase of the phospholipase D type. J Biol Chem 258, 93029306.
  • 105
    Liu J, Bátkai S, Pacher P, Harvey-White J, Wagner JA, Cravatt BF, Gao B & Kunos G (2003) Lipopolysaccharide induces anandamide synthesis in macrophages via CD14/MAPK/phosphoinositide 3-kinase/NF-κB independently of platelet-activating factor. J Biol Chem 278, 4503445039.
  • 106
    Liu J, Wang L, Harvey-White J, Huang BX, Kim HY, Luquet S, Palmiter RD, Krystal G, Rai R, Mahadevan A, et al. (2008) Multiple pathways involved in the biosynthesis of anandamide. Neuropharmacology 54, 17.
  • 107
    Ueda N, Yamanaka K, Terasawa Y & Yamamoto S (1999) An acid amidase hydrolyzing anandamide as an endogenous ligand for cannabinoid receptors. FEBS Lett 454, 267270.
  • 108
    Ueda N, Yamanaka K & Yamamoto S (2001b) Purification and characterization of an acid amidase selective for N-palmitoylethanolamine, a putative endogenous anti-inflammatory substance. J Biol Chem 276, 3555235557.
  • 109
    Tsuboi K, Sun Y-X, Okamoto Y, Araki N, Tonai T & Ueda N (2005) Molecular characterization of N-acylethanolamine-hydrolyzing acid amidase, a novel member of the choloylglycine hydrolase family with structural and functional similarity to acid ceramidase. J Biol Chem 280, 1108211092.
  • 110
    Tsuboi K, Takezaki N & Ueda N (2007a) The N-acylethanolamine-hydrolyzing acid amidase (NAAA). Chem Biodivers 4, 19141925.
  • 111
    Ueda N, Tsuboi K & Uyama T (2010b) N-Acylethanolamine metabolism with special reference to N-acylethanolamine-hydrolyzing acid amidase (NAAA). Prog Lipid Res 49, 299315.
  • 112
    Hong S-B, Li C-M, Rhee H-J, Park J-H, He X, Levy B, Yoo OJ & Schuchman EH (1999) Molecular cloning and characterization of a human cDNA and gene encoding a novel acid ceramidase-like protein. Genomics 62, 232241.
  • 113
    Zhao L-Y, Tsuboi K, Okamoto Y, Nagahata S & Ueda N (2007) Proteolytic activation and glycosylation of N-acylethanolamine-hydrolyzing acid amidase, a lysosomal enzyme involved in the endocannabinoid metabolism. Biochim Biophys Acta 1771, 13971405.
  • 114
    West JM, Zvonok N, Whitten KM, Wood JT & Makriyannis A (2012a) Mass spectrometric characterization of human N-acylethanolamine-hydrolyzing acid amidase. J Proteome Res 11, 972981.
  • 115
    Shtraizent N, Eliyahu E, Park J-H, He X, Shalgi R & Schuchman EH (2008) Autoproteolytic cleavage and activation of human acid ceramidase. J Biol Chem 283, 1125311259.
  • 116
    Armirotti A, Romeo E, Ponzano S, Mengatto L, Dionisi M, Karacsonyi C, Bertozzi F, Garau G, Tarozzo G, Reggiani A, et al. (2012) β-Lactones inhibit N-acylethanolamine acid amidase by S-acylation of the catalytic N-terminal cysteine. ACS Med Chem Lett 3, 422426.
  • 117
    West JM, Zvonok N, Whitten KM, Vadivel SK, Bowman AL & Makriyannis A (2012b) Biochemical and mass spectrometric characterization of human N-acylethanolamine-hydrolyzing acid amidase inhibition. PLoS One 7, e43877.
  • 118
    Sun Y-X, Tsuboi K, Zhao L-Y, Okamoto Y, Lambert DM & Ueda N (2005) Involvement of N-acylethanolamine-hydrolyzing acid amidase in the degradation of anandamide and other N-acylethanolamines in macrophages. Biochim Biophys Acta 1736, 211220.
  • 119
    Tsuboi K, Zhao L-Y, Okamoto Y, Araki N, Ueno M, Sakamoto H & Ueda N (2007b) Predominant expression of lysosomal N-acylethanolamine-hydrolyzing acid amidase in macrophages revealed by immunochemical studies. Biochim Biophys Acta 1771, 623632.
  • 120
    Wang J, Zhao L-Y, Uyama T, Tsuboi K, Wu X-X, Kakehi Y & Ueda N (2008b) Expression and secretion of N-acylethanolamine-hydrolysing acid amidase in human prostate cancer cells. J Biochem 144, 685690.
  • 121
    Tai T, Tsuboi K, Uyama T, Masuda K, Cravatt BF, Houchi H & Ueda N (2012) Endogenous molecules stimulating N-acylethanolamine-hydrolyzing acid amidase (NAAA). ACS Chem Neurosci 3, 379385.
  • 122
    Petrosino S, Iuvone T & Di Marzo V (2010) N-Palmitoyl-ethanolamine: biochemistry and new therapeutic opportunities. Biochimie 92, 724727.
  • 123
    Vandevoorde S, Tsuboi K, Ueda N, Jonsson K-O, Fowler CJ & Lambert DM (2003) Esters, retroesters and retroamide of palmitic acid: pool for the first selective inhibitors of N-palmitoylethanolamine-selective acid amidase. J Med Chem 46, 43734376.
  • 124
    Tsuboi K, Hilligsmann C, Vandevoorde S, Lambert DM & Ueda N (2004) N-Cyclohexanecarbonylpentadecylamine: a selective inhibitor of the acid amidase hydrolysing N-acylethanolamines, as a tool to distinguish acid amidase from fatty acid amide hydrolase. Biochem J 379, 99106.
  • 125
    Solorzano C, Zhu C, Battista N, Astarita G, Lodola A, Rivara S, Mor M, Russo R, Maccarrone M, Antonietti F, et al. (2009) Selective N-acylethanolamine-hydrolyzing acid amidase inhibition reveals a key role for endogenous palmitoylethanolamide in inflammation. Proc Natl Acad Sci USA 106, 2096620971.
  • 126
    Solorzano C, Antonietti F, Duranti A, Tontini A, Rivara S, Lodola A, Vacondio F, Tarzia G, Piomelli D & Mor M (2010) Synthesis and structure–activity relationships of N-(2-oxo-3-oxetanyl)amides as N-acylethanolamine-hydrolyzing acid amidase inhibitors. J Med Chem 53, 57705781.
  • 127
    Saturnino C, Petrosino S, Ligresti A, Palladino C, De Martino G, Bisogno T & Di Marzo V (2010) Synthesis and biological evaluation of new potential inhibitors of N-acylethanolamine hydrolyzing acid amidase. Bioorg Med Chem Lett 20, 12101213.
  • 128
    Duranti A, Tontini A, Antonietti F, Vacondio F, Fioni A, Silva C, Lodola A, Rivara S, Solorzano C, Piomelli D, et al. (2012) N-(2-Oxo-3-oxetanyl)carbamic acid esters as N-acylethanolamine acid amidase inhibitors: synthesis and structure–activity and structure–property relationships. J Med Chem 55, 48244836.
  • 129
    Yamano Y, Tsuboi K, Hozaki Y, Takahashi K, Jin X-H, Ueda N & Wada A (2012) Lipophilic amines as potent inhibitors of N-acylethanolamine-hydrolyzing acid amidase. Bioorg Med Chem 20, 36583665.
  • 130
    Li Y, Yang L, Chen L, Zhu C, Huang R, Zheng X, Qiu Y & Fu J (2012) Design and synthesis of potent N-acylethanolamine-hydrolyzing acid amidase (NAAA) inhibitor as anti-inflammatory compounds. PLoS One 7, e43023.
  • 131
    Prescott SM & Majerus PW (1983) Characterization of 1,2-diacylglycerol hydrolysis in human platelets. Demonstration of an arachidonoyl-monoacylglycerol intermediate. J Biol Chem 258, 764769.
  • 132
    Sugiura T, Kishimoto S, Oka S & Gokoh M (2006) Biochemistry, pharmacology and physiology of 2-arachidonoylglycerol, an endogenous cannabinoid receptor ligand. Prog Lipid Res 45, 405446.
  • 133
    Kano M, Ohno-Shosaku T, Hashimotodani Y, Uchigashima M & Watanabe M (2009) Endocannabinoid-mediated control of synaptic transmission. Physiol Rev 89, 309380.
  • 134
    Fukami K, Inanobe S, Kanemaru K & Nakamura Y (2010) Phospholipase C is a key enzyme regulating intracellular calcium and modulating the phosphoinositide balance. Prog Lipid Res 49, 429437.
  • 135
    Bisogno T, Melck D, De Petrocellis L & Di Marzo V (1999) Phosphatidic acid as the biosynthetic precursor of the endocannabinoid 2-arachidonoylglycerol in intact mouse neuroblastoma cells stimulated with ionomycin. J Neurochem 72, 21132119.
  • 136
    Oka S, Yanagimoto S, Ikeda S, Gokoh M, Kishimoto S, Waku K, Ishima Y & Sugiura T (2005) Evidence for the involvement of the cannabinoid CB2 receptor and its endogenous ligand 2-arachidonoylglycerol in 12-O-tetradecanoylphorbol-13-acetate-induced acute inflammation in mouse ear. J Biol Chem 280, 1848818497.
  • 137
    Ueda H, Kobayashi T, Kishimoto M, Tsutsumi T & Okuyama H (1993) A possible pathway of phosphoinositide metabolism through EDTA-insensitive phospholipase A1 followed by lysophosphoinositide-specific phospholipase C in rat brain. J Neurochem 61, 18741881.
  • 138
    Tsutsumi T, Kobayashi T, Ueda H, Yamauchi E, Watanabe S & Okuyama H (1994) Lysophosphoinositide-specific phospholipase C in rat brain synaptic plasma membranes. Neurochem Res 19, 399406.
  • 139
    Nakane S, Oka S, Arai S, Waku K, Ishima Y, Tokumura A & Sugiura T (2002) 2-Arachidonoyl-sn-glycero-3-phosphate, an arachidonic acid-containing lysophosphatidic acid: occurrence and rapid enzymatic conversion to 2-arachidonoyl-sn-glycerol, a cannabinoid receptor ligand, in rat brain. Arch Biochem Biophys 402, 5158.
  • 140
    Okazaki T, Sagawa N, Okita JR, Bleasdale JE, MacDonald PC & Johnston JM (1981) Diacylglycerol metabolism and arachidonic acid release in human fetal membranes and decidua vera. J Biol Chem 256, 73167321.
  • 141
    Chau LY & Tai HH (1981) Release of arachidonate from diglyceride in human platelets requires the sequential action of a diglyceride lipase and a monoglyceride lipase. Biochem Biophys Res Commun 100, 16881695.
  • 142
    Bisogno T, Howell F, Williams G, Minassi A, Cascio MG, Ligresti A, Matias I, Schiano-Moriello A, Paul P, Williams EJ, et al. (2003) Cloning of the first sn1-DAG lipases points to the spatial and temporal regulation of endocannabinoid signaling in the brain. J Cell Biol 163, 463468.
  • 143
    Jung KM, Astarita G, Zhu C, Wallace M, Mackie K & Piomelli D (2007) A key role for diacylglycerol lipase-alpha in metabotropic glutamate receptor-dependent endocannabinoid mobilization. Mol Pharmacol 72, 612621.
  • 144
    Tanimura A, Yamazaki M, Hashimotodani Y, Uchigashima M, Kawata S, Abe M, Kita Y, Hashimoto K, Shimizu T, Watanabe M, et al. (2010) The endocannabinoid 2-arachidonoylglycerol produced by diacylglycerol lipase α mediates retrograde suppression of synaptic transmission. Neuron 65, 320327.
  • 145
    Gao Y, Vasilyev DV, Goncalves MB, Howell FV, Hobbs C, Reisenberg M, Shen R, Zhang MY, Strassle BW, Lu P, et al. (2010) Loss of retrograde endocannabinoid signaling and reduced adult neurogenesis in diacylglycerol lipase knock-out mice. J Neurosci 30, 20172024.
  • 146
    Hoover HS, Blankman JL, Niessen S & Cravatt BF (2008) Selectivity of inhibitors of endocannabinoid biosynthesis evaluated by activity-based protein profiling. Bioorg Med Chem Lett 18, 58385841.
  • 147
    Bisogno T, Cascio MG, Saha B, Mahadevan A, Urbani P, Minassi A, Appendino G, Saturnino C, Martin B, Razdan R, et al. (2006) Development of the first potent and specific inhibitors of endocannabinoid biosynthesis. Biochim Biophys Acta 1761, 205212.
  • 148
    Bisogno T, Burston JJ, Rai R, Allarà M, Saha B, Mahadevan A, Razdan RK, Wiley JL & Di Marzo V (2009) Synthesis and pharmacological activity of a potent inhibitor of the biosynthesis of the endocannabinoid 2-arachidonoylglycerol. ChemMedChem 4, 946950.
  • 149
    Min R, Testa-Silva G, Heistek TS, Canto CB, Lodder JC, Bisogno T, Di Marzo V, Brussaard AB, Burnashev N & Mansvelder HD (2010) Diacylglycerol lipase is not involved in depolarization-induced suppression of inhibition at unitary inhibitory connections in mouse hippocampus. J Neurosci 30, 27102715.
  • 150
    Fowler CJ (2012) Monoacylglycerol lipase – a target for drug development? Br J Pharmacol 166, 15681585.
  • 151
    Tornqvist H & Belfrage P (1976) Purification and some properties of a monoacylglycerol-hydrolyzing enzyme of rat adipose tissue. J Biol Chem 251, 813819.
  • 152
    Karlsson M, Contreras JA, Hellman U, Tornqvist H & Holm C (1997) cDNA cloning, tissue distribution, and identification of the catalytic triad of monoglyceride lipase. Evolutionary relationship to esterases, lysophospholipases, and haloperoxidases. J Biol Chem 272, 2721827223.
  • 153
    Labar G, Wouters J & Lambert DM (2010a) A review on the monoacylglycerol lipase: at the interface between fat and endocannabinoid signalling. Curr Med Chem 17, 25882607.
  • 154
    Labar G, Bauvois C, Borel F, Ferrer J-L, Wouters J & Lambert DM (2010b) Crystal structure of the human monoacylglycerol lipase, a key actor in endocannabinoid signaling. ChemBioChem 11, 218227.
  • 155
    Bertrand T, Augé F, Houtmann J, Rak A, Vallée F, Mikol V, Berne PF, Michot N, Cheuret D, Hoornaert C, et al. (2010) Structural basis for human monoglyceride lipase inhibition. J Mol Biol 396, 663673.
  • 156
    Dinh TP, Carpenter D, Leslie FM, Freund TF, Katona I, Sensi SL, Kathuria S & Piomelli D (2002) Brain monoglyceride lipase participating in endocannabinoid inactivation. Proc Natl Acad Sci USA 99, 1081910824 (erratum appears in Proc Natl Acad Sci USA 99, 13961).
  • 157
    Tanimura A, Uchigashima M, Yamazaki M, Uesaka N, Mikuni T, Abe M, Hashimoto K, Watanabe M, Sakimura K & Kano M (2012) Synapse type-independent degradation of the endocannabinoid 2-arachidonoylglycerol after retrograde synaptic suppression. Proc Natl Acad Sci USA 109, 1219512200.
  • 158
    Minkkilä A, Saario S & Nevalainen T (2010) Discovery and development of endocannabinoid-hydrolyzing enzyme inhibitors. Curr Top Med Chem 10, 828858.
  • 159
    Long JZ, Li W, Booker L, Burston JJ, Kinsey SG, Schlosburg JE, Pavon FJ, Serrano AM, Selley DE, Parsons LH, et al. (2009b) Selective blockade of 2-arachidonoylglycerol hydrolysis produces cannabinoid behavioral effects. Nat Chem Biol 5, 3744.
  • 160
    Chang JW, Niphakis MJ, Lum KM, Cognetta AB III, Wang C, Matthews ML, Niessen S, Buczynski MW, Parsons LH & Cravatt BF (2012) Highly selective inhibitors of monoacylglycerol lipase bearing a reactive group that is bioisosteric with endocannabinoid substrates. Chem Biol 19, 579588.
  • 161
    Schlosburg JE, Blankman JL, Long JZ, Nomura DK, Pan B, Kinsey SG, Nguyen PT, Ramesh D, Booker L, Burston JJ, et al. (2010) Chronic monoacylglycerol lipase blockade causes functional antagonism of the endocannabinoid system. Nat Neurosci 13, 11131119.
  • 162
    Chanda PK, Gao Y, Mark L, Btesh J, Strassle BW, Lu P, Piesla MJ, Zhang MY, Bingham B, Uveges A, et al. (2010) Monoacylglycerol lipase activity is a critical modulator of the tone and integrity of the endocannabinoid system. Mol Pharmacol 78, 9961003.
  • 163
    Taschler U, Radner FP, Heier C, Schreiber R, Schweiger M, Schoiswohl G, Preiss-Landl K, Jaeger D, Reiter B, Koefeler HC, et al. (2011) Monoglyceride lipase deficiency in mice impairs lipolysis and attenuates diet-induced insulin resistance. J Biol Chem 286, 1746717477.
  • 164
    Nomura DK, Morrison BE, Blankman JL, Long JZ, Kinsey SG, Marcondes MC, Ward AM, Hahn YK, Lichtman AH, Conti B, et al. (2011) Endocannabinoid hydrolysis generates brain prostaglandins that promote neuroinflammation. Science 334, 809813.
  • 165
    Nomura DK, Long JZ, Niessen S, Hoover HS, Ng SW & Cravatt BF (2010) Monoacylglycerol lipase regulates a fatty acid network that promotes cancer pathogenesis. Cell 140, 4961.
  • 166
    Savinainen JR, Saario SM & Laitinen JT (2012) The serine hydrolases MAGL, ABHD6 and ABHD12 as guardians of 2-arachidonoylglycerol signalling through cannabinoid receptors. Acta Physiol (Oxf) 204, 267276 [erratum appears in Acta Physiol (Oxf) 204, 460].
  • 167
    Navia-Paldanius D, Savinainen JR & Laitinen JT (2012) Biochemical and pharmacological characterization of human α/β-hydrolase domain containing 6 (ABHD6) and 12 (ABHD12). J Lipid Res 53, 24132424.
  • 168
    Marrs WR, Blankman JL, Horne EA, Thomazeau A, Lin YH, Coy J, Bodor AL, Muccioli GG, Hu SS, Woodruff G, et al. (2010) The serine hydrolase ABHD6 controls the accumulation and efficacy of 2-AG at cannabinoid receptors. Nat Neurosci 13, 951957.
  • 169
    Marrs WR, Horne EA, Ortega-Gutierrez S, Cisneros JA, Xu C, Lin YH, Muccioli GG, Lopez-Rodriguez ML & Stella N (2011) Dual inhibition of α/β-hydrolase domain 6 and fatty acid amide hydrolase increases endocannabinoid levels in neurons. J Biol Chem 286, 2872328728.
  • 170
    Fiskerstrand T, H'mida-Ben Brahim D, Johansson S, M'zahem A, Haukanes BI, Drouot N, Zimmermann J, Cole AJ, Vedeler C, Bredrup C, et al. (2010) Mutations in ABHD12 cause the neurodegenerative disease PHARC: an inborn error of endocannabinoid metabolism. Am J Hum Genet 87, 410417.
  • 171
    Kozak KR & Marnett LJ (2002) Oxidative metabolism of endocannabinoids. Prostaglandins Leukot Essent Fatty Acids 66, 211220.
  • 172
    Rouzer CA & Marnett LJ (2008) Non-redundant functions of cyclooxygenases: oxygenation of endocannabinoids. J Biol Chem 283, 80658069.
  • 173
    Duggan KC, Hermanson DJ, Musee J, Prusakiewicz JJ, Scheib JL, Carter BD, Banerjee S, Oates JA & Marnett LJ (2011) (R)-Profens are substrate-selective inhibitors of endocannabinoid oxygenation by COX-2. Nat Chem Biol 7, 803809.
  • 174
    Blankman JL, Long JZ, Trauger SA, Siuzdak G & Cravatt BF (2013) ABHD12 controls brain lysophosphatidylserine pathways that are deregulated in a murine model of the neurodegenerative disease PHARC. Proc Natl Acad Sci USA 110, 15001505.