The N-Acylethanolamine-Mediated Regulatory Pathway in Plants

REVIEW

The N-Acylethanolamine-Mediated Regulatory Pathway in Plants

by Aruna Kilarua), Elison B. Blancaflorb), Barney J. Venablesa), Swati Tripathya),Kirankumar S. Mysoreb), and Kent D. Chapman*a)

a) University of North Texas, Department of Biological Sciences, Center for Plant Lipid Research,P. O. Box 305220, Denton, TX 76203-5220, USA

(phone: þ1-940-565-2969; fax: þ1-940-565-4136; e-mail: [email protected])b) Samuel Roberts Noble Foundation, Plant Biology Division, Ardmore, OK 74074, USA

While cannabinoids are secondary metabolites synthesized by just a few plant species, N-acylethanolamines (NAEs) are distributed widely in the plant kingdom, and are recovered inmeasurable, bioactive quantities in many plant-derived products. NAEs in higher plants areethanolamides of fatty acids with acyl-chain lenghts of C12�C18 and zero to three C¼C bonds. Generally,the most-abundant NAEs found in plants and vertebrates are similar, including NAE 16 :0, 18 :1, 18 :2,and 18 :3. Like in animal systems, NAEs are formed in plants from N-acylphosphatidylethanolamines(NAPEs), and they are hydrolyzed by an amidase to yield ethanolamine and free fatty acids (FFA).Recently, a homologue of the mammalian fatty acid amide hydrolase (FAAH-1) was identified inArabidopsis thaliana and several other plant species. Overexpression of Arabidopsis FAAH (AtFAAH)resulted in plants that grew faster, but were more sensitive to biotic and abiotic insults, suggesting that themetabolism of NAEs in plants resides at the balance between growth and responses to environmentalstresses. Similar to animal systems, exogenously applied NAEs have potent and varied effects on plantcells. Recent pharmacological approaches combined with molecular-genetic experiments revealed thatNAEs may act in certain plant tissues via specific membrane-associated proteins or by interacting withphospholipase D-a, although other, direct targets for NAE action in plants are likely to be discovered.Polyunsaturated NAEs can be oxidized via the lipoxygenase pathway in plants, producing an array ofoxylipin products that have received little attention so far. Overall, the conservation of NAE occurrenceand metabolic machinery in plants, coupled with the profound physiological effects of elevating NAEcontent or perturbing endogenous NAEmetabolism, suggest that an NAE-mediated regulatory pathway,sharing similarities with the mammalian endocannabinoid pathway, indeed exists.

1. Occurrence and Distribution of N-Acylethanolamines in Plants and Plant-Derived Products. – N-Acylethanolamines (NAEs)1), the fatty acid ethanolamidesthat are derived from an N-acylphosphatidylethanolamine (NAPE) precursor, areubiquitous in Nature. They have been shown to regulate a variety of physiological

CHEMISTRY & BIODIVERSITY – Vol. 4 (2007) 1933

F 2007 Verlag Helvetica Chimica Acta AG, ZHrich

1) Abbreviations: ABA, abscisic acid; AMI1, amidase 1; AS, amidase signature; AOS, allene oxidesynthase; CB, cannabinoid receptor; COX-2, cyclooxygenase 2; FAAH, fatty acid amide hydrolase;FFA, free fatty acids; HPO, hydroperoxide; LOX, lipoxygenase; MAFP, methylarachidonylflurophosphonate; NAE, N-acylethanolamine, NAPE, N-acylphosphatidylethanolamine; PAL2,phenylalanine ammonia lyase 2; PA, phosphatidic acid; PC, phosphatidylcholine; PE, phosphati-dylethanolamine; PG, phosphatidylglycerol; PS, phosphatidylserine; PLC, phospholipase C; PLD,phospholipase D.

functions in both animals and plants. In this review, we will discuss the occurrence,metabolism, and function of NAEs in plant systems, drawing parallels with animalsystems, where appropriate, and raising the possibility that the role for NAEs as centrallipid mediators in eukaryotic systems is functionally conserved.1.1. N-Acylphosphatidylethanolamines (NAPEs). The biology of NAEs in plants

has its origin in the study of its metabolic precursor, NAPE, which constitutes 1–5% ofthe membrane phospholipids. Early work characterizing the accumulation of NAPEs inresponse to tissue stress in animals was reviewed by Schmid et al. [1]. Although theoccurrence of NAPEs in plants was initially controversial, it was unequivocallydemonstrated by a combination of biochemical and biophysical approaches [2]. NAPEis not abundant under normal physiological conditions. In cotton plants, for example,the NAPE content varies between 1.9 and 3.2 mol-% of the total phospholipids,depending upon the tissue source and developmental stage [3–6]. However, NAPElevels were altered in response to stress [1] [7] [8]. For example, the content of NAPEisolated from potato cells was low in unstressed cells (13�4 nmol/g fresh weight), andincreased up to 13-fold in anoxia-stressed cells, but only when free fatty acids (FFAs)were being released, after ca. 10 h of treatment [9]. TheN-acyl patterns of NAPE weredominated by 18 :1, 18 :2, and 16 :0, but never reflected the FFA composition,suggesting the existence of spatially distinct FFA and phosphatidylethanolamine pools[9]. Similarly, in response to pathogen elicitors such as xylanase, cultures of tobacco-suspension cells showed a fivefold decrease in [14C]-NAPE, and a corresponding sixfoldincrease in [14C]-NAE content [10], suggesting a tight regulation between theendogenous levels of NAPE and NAE.1.2. N-Acylethanolamines (NAEs). NAEs have been known to occur at trace levels

in plants for decades, and were recognized to have anti-inflammatory properties inmammals [11]. However, it was the discovery of the arachidonic acid member of theNAE family (NAE 20 :4, ManandamideN) as an endogenous ligand for the brain CB-1cannabinoid receptor [12] that led to more intensive investigation of this group ofcompounds. Furthermore, the development of analytical techniques permitted accurateidentification and quantification of individual NAE molecular species in animals [13–15] and plants [16–19]. Current methods used for plant tissues are based on Me3Si(TMS) derivatives and gas chromatography/mass spectrometry (GC/MS) single-ionmonitoring as well as isotope-dilution quantification [19].1.3. NAEs in Seeds and Seed Products. 1.3.1. Total NAEs. NAEs have been found in

all seeds examined to date [16] [17] [19] in elevated concentrations relative to adulttissues (Table 1) [20] [21]. NAEs were initially reported in desiccated seeds from eightspecies (pea, soybean, peanut, castor bean, tomato, okra, cotton, and corn), the totalNAE concentrations ranging from 490 (pea) to 1,608 (cotton cv. Stonville 7A glandless)ng/g fresh weight [16]. This study demonstrated an approximate threefold variationamong the species examined. When nine cultivars of cotton were examined, theyexhibited a range of total NAE content similar to that seen among the eight distinctspecies [16].In a subsequent study, legumes were surveyed for NAE content [19]. The legume

family is the third-largest flowering-plant family, with more than 700 genera and some20,000 species, representing a wide range of morphological and ecological/life historycharacteristics. A total of 14 taxa were selected, with twelve species representing ten

CHEMISTRY & BIODIVERSITY – Vol. 4 (2007)1934


Table1.RangeofConcentrations(in

mg/gfreshweight)ofN-Acylethanolamine(NAE)SpeciesinSelectedPlantsandPlant-DerivedProducts.

Abbreviations:OE¼FAAHoverexpressor;KO¼FAAHknockout.

Tissue

Taxona )

NAESpecies

Ref.

Total

12:0

14:0

16:0

18:0

18:1

18:2

18:3

Seeds

Tomato

0.75

0.15

0.03

0.10

0.03

0.13

0.29

n.d.b)

[7]

Castor

0.62

0.14

0.04

0.05

0.02

0.08

0.25

n.d.

Okra

0.76

0.15

0.04

0.13

0.03

0.12

0.29

n.d.

Corn

1.1

0.18

0.02

0.20

0.07

0.33

0.38

n.d.

Cotton

0.75–2.3

0.15

0.06

0.35

0.06

0.16

0.87

n.d.

Anacachoorchidtree

0.31

0.03

0.01

0.04

<0.01

0.14

0.07

0.02

[19]

Texasbluebonnet

1.5

0.01

0.01

0.37

0.06

0.74

0.81

0.10

Peanut

17<0.01

0.02

3.73

0.69

7.96

4.54

0.12

Alfalfa

7.4

<0.01

<0.01

1.1

0.47

1.0

2.0

2.8

Barrelclover

45<0.01

0.37

132.0

9.6

128

Gardenpea

0.58

<0.01

<0.01

0.10

0.04

0.22

0.17

0.04

Soybean

320.46

0.51

6.7

1.6

4.9

133.5

Seeds

Arabidopsisthaliana(wildtype)

2.0

0.07

0.03

0.06

0.03

0.20

1.3

0.24

[20]

Arabidopsisthaliana(KO)

2.7

0.04

0.02

0.14

0.03

0.4

1.7

0.40

Arabidopsisthaliana(OE)

1.0

0.06

0.02

0.02

0.02

0.10

0.62

0.12

Seedlings


0.22

0.06

0.02

0.02

0.02

0.02

0.06

0.03

[20]


0.26

0.08

0.03

0.01

0.02

0.01

0.09

0.03

Arabidopsisthaliana(OE)

0.20

0.07

0.03

0.03

0.02

0.01

0.04

0.01

Adultvegetative


0.05

0.01

<0.01

<0.01

<0.01

<0.01

0.02

0.01

n.p.c )


0.18

0.02

<0.01

0.01

0.01

0.01

0.07

0.05

Arabidopsisthaliana(OE11)

0.05

0.01

<0.01

<0.01

<0.01

<0.01

0.01

0.01

Plant-derived

d)

Variousfoods

––

––

–�1.1

�2.8

–[21]

Cottonseed

(MfinishedmealN)

8.8

0.08

0.07

2.6

0.20

1.4

4.5

–[17]

Soylecithin

>2,000

––

950

––

1,200

–n.p.c )

a )Tomato,Lycopersiconesculentum;castor,Ricinuscommunis;okra,Abelmoschusesculentus;corn,Zeamays;cotton,Gossypium

hirsutum;anacacho

orchidtree,Bauhiniacongesta;Texasbluebonnet,Lupinustexensis;peanut,Arachishypogaea;alfalfa,Medicagosativa;barrelclover,Medicagotruncatula

cv.Jemalong;gardenpea,Pisum

sativum

cv.Taos;Soybean,Glycinemaxcv.Dare.

b)Notdetermined.c )Notpublished(CenterforPlantLipidResearch,

Denton,TX,U.S.A.).d)Onlythemost-abundantNAEswerequantifiedorreported.

genera in four major sub-families. In this study, the total NAE content in desiccatedseeds ranged over three orders of magnitude from 170 (kidney bean) to 44,600 (Barrelclover cv. Jemalong) ng/g. While there was a general tendency for seeds with higherlipid content to have higher NAE concentrations, even lipid-normalized NAEconcentrations varied over two orders of magnitude from 3,010 (Anacacho orchidtree) to 350,000 (Barrel clover cv. Jemalong) ng/g of lipid. Although the most-primitivelegume taxa tended to be among the lowest in NAE content, a strong phylogeneticcorrelation with NAE concentration was not observed. Thus, while NAEs have beendetected in all seeds examined, their relative contribution to both fresh and lipid massappears to be highly taxon-specific, even to the cultivar level, and reasons for theseobserved differences remain an intriguing and unresolved aspect of understanding thebiology of NAEs in plant seeds.1.3.2. Patterns of Dominance in Molecular Species in Seeds. The relative dominance

of molecular species contributing to the total NAE content in seeds parallels thedistribution characteristic of acyl groups in plant membrane lipids in general, and inNAPEs specifically [16] [19] [22]. Thus, NAEs are dominated by saturated C12, C14, C16,and saturated or unsaturated C18 acyl chains, the latter comprising one to three C¼Cbonds. The NAEs with C16 and C18 acyl chains typically are the most abundant, withminor contributions from C12 and C14. As in animal systems, these minor NAEs havepotent biological activity. Among the C18 species, NAE 18 :2 is typically the mostabundant. Exceptions were seen in peanut, in which NAE 18 :1 exceeded NAE 18 :2,and in alfalfa, in which NAE 18 :3 exceeded NAE 18 :2; these changes mirror therelative abundance of the corresponding acyl groups in these species [19].1.3.3. NAEs in Seed Products. The dominant NAEs found in seeds, the unsaturated

C18 species and NAE 16 :0, are known to have biological activity in mammaliansystems. As mentioned previously, the anti-inflammatory activity of NAE 16 :0 hasbeen recognized for many years [11] [23] [24]; it appears to involve a mechanism thatdownregulates fatty acid amide hydrolase (FAAH) activity via the peroxisomeproliferator-activated receptor-a [25]. NAE 18 :2 has been shown to compete withendogenous NAEs for FAAH [26]. Therefore, the two predominant plant NAE speciesmay interact with each other to potentiate activity of CB-receptor agonists byinterfering with both the production and activity of the principle enzyme responsiblefor agonist breakdown. The therapeutic potential of NAEs has led to a search forreadily accessible, inexpensive sources of natural products in which they are enriched.Because of the high commercial production volume of cottonseed, oil-mill fractions

were examined for NAE content [17]. There was a single clearly enriched fraction inthe milling process. The Mfinished mealN fraction (Table 1), which is a product resultingfrom the fortification of hexane-extracted cottonseed with the soap stock fraction, hadan approximately eightfold increased NAE concentration, when compared todesiccated seed. However, this enriched fraction still had NAE concentrations farbelow those observed in the most-elevated taxa represented in the previously describedlegume survey [19]. Soybeans, another high commercial-production-volume oil seed,may lose most of their NAE content during the de-gumming process [22], as the crudesoy-lecithin fraction seems to be especially enriched, whereas refined soybean oilcontains little or no NAEs. In a survey of eight commercial soy-lecithin products, thetotal NAE content was found to vary widely; however, four of the products had total


NAE concentrations in the low milligram-per-gram range (B. J. Venables, K. D.Chapman, unpublished data). Lecithins appear to represent the richest natural sourceyet identified for this family of compounds.

2. NAE Metabolism in Plants. – NAE Metabolism is highly conserved in animalsand plants, although there are some notable differences (Fig. 1). It has been proposedthat the saturated and unsaturated species of NAEs are formed from theircorresponding N-acylphosphatidylethanolamine, a minor membrane component, in asingle enzymatic step catalyzed by a phospholipase D (PLD) [8] [27] [28]. NAEs arefurther catabolized either by hydrolysis, by a fatty acid amide hydrolase to FFAs andethanolamines [29–32], or, alternatively, some polyunsaturated NAEs are oxidized bylipoxygenase (LOX) and allene oxide synthase (AOS) to yield NAE oxylipins[33] [34]. In plants, FFAs formed from the hydrolysis of NAE or glycerophospholipidscan be incorporated directly into the N-position of NAPE by NAPE synthase [2] [35].Progress made in understanding NAE metabolism in plants is discussed further.

2.1. NAE Precursor – NAPE Biosynthesis. N-Acylphosphatidylethanolamine is anunusual phospholipid, with a third acyl moiety linked to the ethanolamine head groupof phosphatidylethanolamine (PE), and a minor constituent of the membranes ofplants, animals, and some microorganisms [1] [2]. In plants, NAPE synthase wasinitially purified from the cotyledons of imbibed cottonseeds, and was discovered to bea membrane-bound enzyme synthesizing NAPE by direct acylation of PE with non-esterified FFAs [2] [35] [36]. However, in animals, NAPE is synthesized by atransacylation reaction in which the sn-1-O-acyl moiety of phosphatidylcholine (PC)

Fig. 1. The Metabolism of N-Acylethanolamine (NAE) is conserved in eukaryotic organisms. In bothplants and animals, NAEs are formed by the hydrolysis of the membrane component N-acylphospha-tidylethanolamine (NAPE) by a phospholipase D (PLD). NAEs are hydrolyzed by fatty acid amidehydrolase (FAAH) into ethanolamine and free fatty acids (FFA). Polyunsaturated NAEs can also beoxidized to NAE oxylipins via the lipoxygenase (LOX) or cyclooxygenase (COX) pathway in plants andanimals, resp. NAPE is formed from FFA via NAPE synthase in plants, and by coordinatedacyltransferase-transacylase in animals. Similarities and differences in the metabolic pathways between

animals and plants are color-coded.


is transferred to the N-atom of PE without an FFA intermediate (Fig. 1) [37].Furthermore, NAPE synthase in plants incorporates acyl moieties from acyl CoA orPC-100 to PC-1000 less efficiently than FFAs, and they show ATP-, Ca2þ-, and CoA-independent acyltransferase activity [2]. These unusual biochemical properties suggestthat in vivo NAPE synthase may scavenge FFAs for rapid recycling of fatty acids backinto membrane-associated NAPE, as noted by in vivo accumulation of FFAs followedby rapid increase in NAPE levels in potato cells under hypoxic stress [9] [16].In both animals and plants, NAPE is hydrolyzed by PLD (discussed below), and the

types of NAEs formed are usually the reflection of the acyl groups present in the NAPEprecursor pool [8]. This suggests that the NAPE precursors determine the nature of theNAE pool, leaving little room for influence by the molecular selectivity of NAPE bythe NAPE-PLDs [38] [39]. Molecular characterization of NAPE synthase will furtherelucidate the functional role(s) of NAPEs in plants and animals in terms of signaltransduction or cellular protection.2.2.NAE Formation.Amembrane-bound phosphodiesterase of the phospholipase-

D type has been initially characterized in rat-heart and rat-brain microsomes [40] [41].However, the recent molecular characterization of this Ca2þ-activated NAPE-hydro-lyzing PLD inmouse (NAPE-PLD) [38] has facilitated several advances in decipheringNAE formation in animals. NAPE-PLD was identified to be a member of the Znmetallo-lactamase family, and is highly conserved in mammals. This membrane proteinis structurally and catalytically distinguishable from all other known enzymes and PLDs[38] [39], and its expression is age- and tissue-dependent [42]. Interestingly, the yeastgenome was also shown to contain one gene (YPL103c), with 50.3% homology tohNAPE-PLD [43]. However, no such homology to mammalian NAPE-specific PLD inplants has been reported [44].Our knowledge with regard to the regulation of NAEmetabolism in plants is still in

its early stages. Despite the identification of multiple PLD genes in plants (twelve inArabidopsis [45]), a PLD that is exclusively responsible for hydrolysis of NAPE toNAE and phosphatidic acid (PA) has not been identified. Instead, in vitro studiesshowed that recombinant plant PLD-b and -g isoforms, expressed in E. coli, couldcatalyze the formation of NAE from NAPE [46]. Unlike the newly identifiedmammalian NAPE-PLD, which was inactive with PC and PE [38] [40] [41] [47], bothPLD-b and -g were able to hydrolyze PE, PC, phosphatidylglycerol (PG), andphosphatidylserine (PS), in addition to NAPE [46]. Thus, in plants, PLD-b and -g, twopolyphosphoinositide-dependent PLDs [46], are capable of forming NAE; however,work remains to unequivocally identify the in vivo mechanism for NAE formation.Overexpression of mouse NAPE-PLD resulted in a 50–90% decrease in NAPE,

and in a 1.5-fold increase in the total amount of NAEs, suggesting that the recombinantNAPE-PLD utilizes endogenous NAPE as a substrate in the cell [39]. Conversely,targeted disruption of NAPE-PLD gene in mouse resulted in lower levels of saturatedNAEs, but polyunsaturated NAE levels remained essentially unchanged, suggestingthat multiple andmechanistically distinct routes for NAE biosynthesis exist that may beNAE species- and organ/tissue-specific [48]. In support of this concept, the existence ofan uncharacterized a/b-hydrolase 4 (Abh4), a (lyso) NAPE-lipase [49], and aphospholipase C (PLC), inducible by bacterial endotoxins [50], were reported toparticipate in an NAPE-PLD-independent pathway(s) for NAE biosynthesis. Although


in vivo evidence for NAPE-PLD, or an alternate enzyme catalyzing the formation ofNAE in plants, is lacking, both in vitro and in vivo radiolabeling experiments stronglysuggest that NAPE is, indeed, the precursor for NAEs in plant cells [16] and, like inanimals, plants may also possess multiple ways to synthesize NAEs.2.3. NAE Hydrolysis. In vertebrates, the endocannabinoid signaling pathway that

regulates diverse physiological processes is principally terminated by FAAH, whichhydrolyzes NAEs into their corresponding FFAs and ethanolamine [30]. FAAH is amember of the amidase superfamily of proteins, which all contain a conserved amidase-signature (AS) sequence, and the crystal structure of rat FAAH has revealed importantinsights into its structure and function [30]. A bioinformatics approach led to themolecular identification of an FAAH homolog in Arabidopsis (At5g64440) [31].Recombinant FAAH proteins from plant and animal sources, expressed in E. coli,exhibited similar biochemical properties, including similar affinities for a broad rangeof NAEs and potent inhibition by the active-site-directed enzyme inhibitor methyl-arachidonyl fluorophosphonate (MAFP) [31] [51]. One notable difference was thecomparative insensitivity of the plant FAAH enzymes to the mammalian FAAHinhibitor URB597 [52].The gene architecture was compared for three plant FAAH homologues, including

a legume (M. truncatula), a monocot (rice), and a non-legume dicot (Arabidopsis)[52]. Although relatively complex, the genes were highly conserved in terms of overallexon organization; each plant species had an FAAH gene with 19 or more exons, andsome variability in the size and number of the two or three exons encoding the 5’-UTR.The cDNAs corresponding to these FAAH genes enabled the identification of threespecific protein domains spanning the AS sequence, as well as three other domains ofunknown function near the C-terminus of the plant FAAH proteins [52]. Two of theseunknown domains in plants shared some similarity with rat FAAH [22]. Five aminoacid residues, determined to be important for catalysis by rat FAAH, were absolutelyconserved within the FAAH sequences of six plant species. Homology modeling, usingthe rat FAAH crystal structure as a template, predicted a conserved active-site core inplant FAAHs (Fig. 2) [52]. In all, while there was only 20% amino acid sequenceidentity over the entire length of mammalian and plant FAAH proteins, there was ahigh degree of homology within distinct domains, and considerable overlap in catalyticproperties.Recent studies with FAAH-deficient mice revealed higher levels of NAE 20 :4,

NAE 16 :0, and NAE 18 :1 in their tissues compared to wild-type mice, supporting arole for FAAH in NAE metabolism [53] [54]. Similarly,Arabidopsis seeds with alteredFAAH expression showed altered endogenous levels of NAEs [55]. NAE Levels inseeds of FAAH insertional mutants were ca. 30% higher than in wild-type seeds, and inFAAH overexpressors they were ca. 50% lower, with most of the differences found inthe saturated C16 and unsaturated C18 species. Furthermore, early developmentaldeclines in NAE content have been correlated with increased FAAH activity inseedlings [33] [55]. NAE Levels in 8-d-old seedlings showed similar but less-pronounced differences in NAE content in FAAH-altered lines, with 10% higheramounts in T-DNA knockout seedlings and 15% lower amounts in FAAH over-expressors. Seedlings of FAAH-altered plants showed anticipated differences insensitivity toward exogenous NAEs [55] (see also Sect. 4). The tolerant AtFAAH


overexpressors and hypersensitive T-DNA insertional mutants established therelevance of AtFAAH in vivo for the metabolic inactivation of NAE in plants.Perhaps just as important, the depletion of NAE levels in FAAH T-DNA-disruptedseedlings indicates that an alternate pathway for NAE metabolism may exist inseedlings for the depletion of endogenous NAEs.


Fig. 2. Modeling of plant FAAH proteins. a) An alignment of the conserved amidase-signature (AS)sequence of rat, Arabidopsis, Medicago, and rice FAAH proteins. b) Domain architecture of plantFAAH proteins was proposed based on the consensus sequence. The catalytic-triad residues K142, S217,and S241 of the rat sequence are highly conserved across the plant FAAH proteins. c) Homology-basedmodeling of all plant FAAH domains, using the rat FAAH protein, with the MAFP inhibitor docked inthe active site. d) Overlay of catalytic residues in the plant FAAH active sites, showing nearly directsuperposition of each side chain. Side chains of all plant FAAH models are shown, and amino acidpositions are numbered according to theArabidopsis sequence. The sequences in a), b), and c) are color-coded to show their position in the domain architecture and homology modeling (parts of this figure

were taken from [52], reprinted with kind permission from Biochimica Biophysica Acta).

The possibility for the presence of alternate routes for NAE metabolism inmammals is circumstantially supported by pharmacological studies in FAAH(� /� ) mice[56], and led to the discovery of alternate enzymes that hydrolyze NAE, such as NAEacid amidase (NAAA) [57] [58], a second mammalian FAAH (FAAH-2) [32], andidentification of alternate NAE derivatives [59]. Plant homologs of NAAA were notfound; however, plant homologs of ceramidase [60] need to be tested for NAE-hydrolyzing activity. Mammalian FAAH-2 (FLJ31204), revealed by a functionalproteomic analysis, displayed overlapping, but distinct, tissue-distribution, substrate-selectivity, and inhibitor-sensitivity profiles compared with the original FAAH enzyme[32]. Interestingly, the FAAH-2 gene is present in primates, as well as in a variety ofdistantly related vertebrates, but not in murids [32], and no close homology in plantswas identified; however, several amidases in plants are candidates for NAE hydro-lases.In silico analysis of the seven AS proteins in the Arabidopsis genome placed these

proteins in two sub-groups and two single outlying proteins [61]. Of these, onlyAtFAAH and amidase 1 (AMI1) have been characterized; while AtFAAH metabo-lized NAEs and oleamide, and AMI1 was specific for indole-3-acetamide and 1-naphthaleneacetamide [61]. Here, construction of a broader phylogenetic tree [62] thatincludes amidases from Arabidopsis, Medicago, rice (Oryza), human, rat, and yeastclusters the proteins into five subgroups (Fig. 3). Interestingly, plant FAAH wasgrouped together (group III), but was separated from the mammalian FAAH-1 andyeast amidase (group I). The recently characterized hFAAH-2 diverged considerablyfrom hFAAH-1 (group I), and both of these proteins hydrolyze NAEs. This suggeststhat other uncharacterized amidases in plants may be discovered that have NAEhydrolase activity, and the tree shown in Fig. 3 points to some candidates for testingsuch as Arabidopsis NP196353 (group II), or Medicago ABE86169, and riceABA95925 (group III).The amidases annotated as Mglutamine-tRNA-synthetase-likeN were grouped

together and included human amidase, NP060762 (group IV). Finally, the recentlycharacterized AMI1 and a chloroplast outer-membrane protein (TOC64-like) inArabidopsis, as well as two uncharacterized amidases from rice, were clustered in groupV. Group II had three proteins branching directly from the origin; they are yet to becharacterized. Putative FAAH genes were identified for additional plant species byconceptual translation of their mRNA sequences and identifying homology with theAtFAAH (Table 2). Identification and characterization of additional plant amidaseswill elucidate potential alternate enzymes that influence the metabolic fate of NAEs inplants.2.4. NAE Oxidation. In plants, polyunsaturated fatty acids commonly undergo

oxygenation via the lipoxygenase (LOX)-mediated pathway to form various oxylipins,many of which are important metabolites in plant physiology [63] [64]. Similarly,polyunsaturated NAEs, anandamide [65], N-linoleoylethanolamine [66], and a/g-N-linolenoylethanolamine (NAE 18 :3) [34] are suitable substrates for plant lipoxyge-nases. Kinetics studies revealed that the lipoxygenase products, the hydroperoxides ofNAE 18 :2 and NAE 18 :3, can be converted readily (or even better than thecorresponding FFAs) by hydroperoxide lyase (HPO lyase) and/or allene-oxidesynthase to yield NAE-oxylipins [34].


Experiments with imbibed cotton seeds indicated that NAE 18 :2 was metabolizedin a stereoselective manner by 13-LOX activity, followed by 13-AOS, giving rise to anovel oxylipin-12-oxo-13 hydroxy-octadecenoylethanolamide [33]. The fate of NAE-derived oxylipins in plants is unclear. In animal systems, cyclooxygenase 2 (COX-2)acts on anandamide, which yields the prostaglandin analog PGE2-ethanolamide [67].These eicosanoid ethanolamides are important signaling compounds that participate indiverse physiological processes [68]. NAEOxylipins are yet to be discovered to play animportant mediator role in plant biology. In fact, the rapid depletion of NAEs during


Fig. 3. Other Amidases in Plants. The amidase-signature (AS) sequence (IPR000120) was used toidentify putative amidase proteins in three plant species,Arabidopsis thaliana (AT),Oryza sativa (rice),and Medicago truncatula (MT). Multiple amidase sequences from the three species, along with thecharacterized FAAH sequences from human, rat, and yeast, were aligned using ClustalW, with defaultparameters. A radial phylogram was generated using the TreeView software (see http://taxonomy.zoo-logy.gla.ac.uk/rod/treeview.html). The amidases, candidate and known, were clustered into five groups;human, rat, and yeast FAAH in group I, and the characterized plant FAAHs in group III. The recentlyidentified Arabidopsis (AT-AMI1) that hydrolyzes auxin amides is in group V; the glutamyl-tRNAamidotranferases are in group IV. Group II contains several candidate amidases that have yet to becharacterized. The branch length indicates the relative observed divergence among the polypeptides. The

numbers in parentheses are sequence identifiers.

seed germination and seedling development, especially in the AtFAAH knockout linesof Arabidopsis (AtFAAH) [55], may suggest that NAE oxidation is important forgermination and seedling development. Future advances will require the accurateprofiling of NAE oxylipins in plant systems to better understand their functionalsignificance.


Table 2. Plant Sequences from the UniGene Database Showing <30% Alignment (ProtEST) with N-Acylethanolamine Hydrolase from Arabidopsis thaliana (607 amino acids)

UniGeneEntry

Generic name GenBankAccession

Alignmentidentity [%]

Alignmentlengtha)

Proteincoordinates

Nucleotidecoordinates

At.19619 Arabidopsis AY308736 100 607 1–607 155–1975Gma.10626 Glycine BE348179 78.8 52 537–588 588–743Gma.16265 Glycine BQ612594 68.9 212 301–512 2–637Gma.22765 Glycine BE823508 33 67 538–604 179–493Gma.26030 Glycine BE021799 59 83 343–425 437–685Hv.1711 Hordeum CB880279 60 54 551–604 408–572Hv.18023 Hordeum BF624258 35 134 5–138 81–494Hv.19588 Hordeum AJ467997 44 147 459–605 15–458Hv.6024 Hordeum BI960205 57.8 44 563–606 439–573Hv.7973 Hordeum AW983509 52.2 232 331–562 1–693Les.13527 Lycopersicon BI929799 85 41 269–309 1–123Les.1465 Lycopersicon 132131F 63.6 607 1–607 49–1878Les.7200 Lycopersicon AW223874 38 206 323–528 2–607Les.7279 Lycopersicon BI924944 60 191 1–191 135–707Les.905 Lycopersicon 132142R 65.2 253 113–365 735–1493Lsa.8191 Lotus BQ874500 39 239 303–541 1–696Mtr.2133 Medicago CA921451 48.1 182 426–607 249–794Mtr.6927 Medicago BF646027 37 212 20–231 9–641Os.14121 Oryza 002–116-C08 43.7 603 5–607 113–1912Os.52332 Oryza J013066H12 58 50 217–266 265–414Os.9707 Oryza J033061L19 60.1 535 71–605 334–1941Ppa.2800 Physcomitrella BJ608857 51.1 94 512–605 449–730Ppa.3177 Physcomitrella BJ195266 40.4 99 197–295 315–611Ppa.320 Physcomitrella BJ590804 44.7 206 401–606 121–741Ppa.3832 Physcomitrella BJ173862 52.1 141 464–604 206–631Pta.14954 Pinus CF672587 33.3 200 5–204 103–720Pta.17534 Pinus CO176265 62 263 84–346 2–790Stu.1236 Solanum CV502261 63.1 240 368–607 5–727Stu.14712 Solanum BQ047488 36.8 106 371–476 424–657Stu.6615 Solanum CV474916 69.1 122 483–604 18–386Ta.1047 Triticum CK161883 42.5 281 5–285 95–949Ta.9538 Triticum CD930995 51.8 253 338–590 6–761Vvi.12041 Vitis CF513783 70.8 153 452–604 350–811Vvi.13519 Vitis CN547490 47.6 144 464–607 234–668Zm.26498 Zea AY110504 60.5 123 482–604 217–588Zm.7100 Zea AY106197 38.7 212 336–547 23–637

a) In terms of amino acid residues.

3. Biochemical Targets of NAEs in Plants. – NAEs are likely to mediate theircellular effects in plants through the direct interaction with protein targets. But like inanimal systems, it seems that there are multiple targets that contribute to thepleiotropic effects of NAEs in plants (see below). Evidence at this stage points to twobiochemical targets: a membrane-bound protein (NAE-binding protein), similar to theCB-receptor paradigm of neuronal systems, and a phospholipase (PLD-a) withstructural features unique to plant systems. Certainly, more targets are likely to bediscovered in the future, and opportunities abound to identify the components of thesepathways and to understand how their interactions influence plant growth andresponses to cellular stressors.3.1. NAE-Binding Protein in Plants. The cannabinoid receptors, CB1 and CB2, and

the vanilloid-receptor ion channels (e.g., TRPV1) are cellular binding sites for NAE20 :4 (anandamide), 2-arachidonylglycerol, and other (yet unidentified) endogenouscannabinoids [69] [70]. Several synthetic cannabinoids (WIN 55, 212-2, AM 281,SR144528, etc.) for each CB-receptor subtype have been developed to characterizetheir structure–activity relationships (SAR) [71]. In studies aimed at identifying NAEreceptors in plants, the CB-receptor-subtype antagonists AM281 and SR144528 wereused for elucidating possible components of the NAE signal-perception and -trans-duction events. Both CB-receptor antagonists effectively suppressed the characteristicNAE-induced, defense-related activities in tobacco, the alkalization of the extracellularspace, and the induction of phenylalanine ammonia lyase (PAL2) mRNA accumulation[72]. Consequently, a ligand-binding assay for the CB receptor [73] was modified forassessing binding of [3H]NAE 14 :0 directly to tobacco-suspension cells. Identificationof a saturable, high-affinity binding activity (Kd¼74 nm) for [3H]NAE 14 :0 was thefirst indication of the occurrence of a binding protein for NAE in plants that mightmediate its physiological actions [72]. The NAE-binding affinity was comparable to Kdvalues of not only some pathogen elicitor proteins such as elicitins (Kd¼5.8–13.5 nm)[74] [75]) and harpin (Kd¼425 nm) [76], but also of the Kd values of CB-receptoragonists and antagonists in mammalian systems [77].The partial CB-receptor agonist [3H]NAE 20 :4 [78] did not bind to tobacco cells,

whereas the synthetic CB-receptor antagonists AM281 and SR144528 did compete for[3H]NAE 14 :0 binding. Additionally, the binding activity of [3H]NAE 14 :0 wascompletely eliminated by higher concentrations (1.0 mm) of both CB-receptor-subtypeantagonists, suggesting that the NAE-binding protein in tobacco cells might share somebiochemical similarities with mammalian CB receptors [72]. The NAE-binding activitywas characterized in isolated tobacco cell membranes, and in membranes from leaves ofseveral plant species, including Arabidopsis. Moreover, this binding protein wassolubilized in an active form using the non-ionic detergent dodecylmaltoside [72] [79].However, conventional protein-purification strategies have yet to reveal the identity ofthis NAE-binding protein, and no obvious homologs of the CB receptors are evident inthe Arabidopsis genome.The key to obtaining further support for NAE receptors in plants will require the

molecular identification of proteins with NAE-binding properties, and evidence inplanta for their involvement in NAE-regulation of physiological responses. It is possiblethat, as the functional domains of mammalian NAE-binding proteins are bettercharacterized, a functional-genomics approach may help to identify candidates for


membrane-bound targets of NAEs in plant systems. A recent micro-array analysis ofRNA isolated fromArabidopsis leaves revealed that 8% of the 286 genes, up-regulatedby NAE 14 :0 treatment, were defense-related, and their expression was also reversedby the CB-receptor antagonist SR144528 (S. Tripathy ; unpublished results). Overall, itseems likely that a membrane receptor for NAEs in plants accounts for at least part ofNAE-mediated activity in vivo, and this remains an attractive hypothesis for NAEaction in plants.3.2. Phospholipase D-a (PLD-a). Several years ago, a survey of Arabidopsis PLD

isoforms indicated that the major plant PLD, i.e., PLD-a, was inactive toward NAPEs[46]. Interestingly, NAEs were found to be potent inhibitors of PLD activity towardother phospholipid substrates, and this inhibition was most potent (nanomolarconcentrations in vitro) with short- to medium-chain saturated NAEs [80]. Theinhibition of PLD-a activity by NAE was not seen for other plant PLD isoforms, or forthe well-characterized PLD from Streptomyces, which raises the possibility that NAEinhibition may be a mechanism for the selective regulation of PLD-a activity in planta.Indeed, the administration of NAE to plant tissues phenocopied the responses in PLD-a-antisense-suppressed plants [45] [81] in terms of regulating stomatal aperture [80].And now, NAE 12 :0 is becoming more widely used as a PLD-a inhibitor in plant-cellphysiology (see below) [82]. Thus, one direct target in plants that mediate NAE actionis likely to be PLD-a.PLD-a is involved in a number of stress-signaling pathways in plants, and its

interaction with abscisic acid (ABA) signaling has been established. PLD-a is involvedin the activation of ABI1, a negative regulator of ABA responses, and the activity ofPLD-a appears to be modulated by its direct interaction with the G-protein GPA1 [83–85]. It is possible that NAE interferes with the interaction of PLD-a with GPA1, andmodulates its enzymatic activity, thereby modulating downstream ABA-responsivepathways. Further work is required to determine the precise interaction of NAE andPLD-a in planta and the cellular consequences of this action, but some concepts areexplored further below (see Sect. 4.2 and 4.2.2). Nonetheless, models of an NAEregulatory pathway in plants must take into account the regulation of PLD-a.

4. Function of NAE Metabolism in Plant Growth and Stress Responses. – Theevidence for the occurrence of an NAE-metabolic machinery in plants and in vitrobiochemical studies points to specific plant cellular targets [31] [72] [80]. However,unlike research in animal systems, which is facilitated by well-established assays foranalyzing anandamide functions in vivo [30] [86], NAE research in plants has beenhindered by the lack of well-defined cellular and physiological parameters for assessingNAE action in planta. In attempts to overcome these deficiencies, it was recently shownthat exogenously applied NAEs have potent growth-inhibitory effects during earlydevelopment of Arabidopsis seedlings (Fig. 4,a) [82] [87], which presents an easilyquantifiable assay to assess how alteration of specific enzymes involved in NAEmetabolism might influence plant development (see, e.g., [55]) and responses to bioticand abiotic stresses.4.1. NAEs Negatively Regulate Seedling Growth and Development. NAEs are

comparatively abundant in desiccated seeds, and their levels are depleted during seedgermination, indicating that the catabolism of NAEs might be a requisite for early


seedling establishment [7] [33] [55]. Decreased NAE levels after 4 h of inhibition werereported for peanut, pea, and cotton [7], and at 24–48 and 96–192 h post germinationfor Arabidopsis [55]. For A. thaliana, the levels declined from ca. 2,500 ng/g indesiccated seeds to <1,000 ng/g at 24–48 h, and to ca. 500 ng/g at 96–124 h,respectively [55], and continued to decline another order of magnitude to ca. 50 ng/gin adult leaves (V. Vadapalli, C. A. Waggoner, B. J. Venables, K. D. Chapman,unpublished results).A. thaliana seeds germinated and grown in the presence of micromolar concen-

trations of exogenous NAE 12 :0 exhibited reduced growth, as measured by organ sizesof seedlings [55] [87]. Reductions in root length, impairment of root-hair development,swelling of root tips, reduction in the length of hypocotyls, and reduction in the surfacearea of cotyledon were all reported to be dependent upon NAE concentration, andwere not observed with FFA analogs or NAE 16 :0 [87]. These growth parameters weremore profoundly reduced in AtFAAH knockouts (T-DNA insertional mutants) grownin the presence of NAE 12 :0, whereas seedling growth of AtFAAH overexpressorgrowth was much less affected by exogenous NAE, indicating that NAE-mediatedinhibition of seedling growth can be mitigated by upregulation of AtFAAH [55].


Fig. 4. NAE Effects on Growth and Cytoskeletal Organization in Arabidopsis seedlings. a) Arabidopsisseedlings, grown in the presence of 30 mm NAE 12 :0 for 5 d, display a dramatic reduction in size. Theinhibitory effects of NAE 12 :0 on seedling growth is manifested at the cellular level. b) Primary roots of4-d-old seedlings germinated in the presence of 50 mm NAE 12 :0 are characterized by swollen cells alongthe elongation zone. This is in contrast to the cylindrical shape of the cells in untreated seedlings. c)Cortical microtubules in untreated seedlings are oriented perpendicular to the long axis of the cell,

whereas roots treated with NAE 12 :0 exhibit randomly oriented microtubules.

AtFAAH Overexpressors, with lower levels of endogenous NAEs in seeds, showedenhanced growth phenotypes in seedlings compared with wild-type and AtFAAHknockouts under normal conditions. AtFAAH overexpressors showed quantifiableincreases in root length, hypocotyl length, cotyledon size, petiole length, and leaf size[55], suggesting a clear relationship between FAAH-mediated NAE metabolism andnormal seedling growth and development.Some NAE types may be of particular importance for membrane protection during

desiccation/rehydration stress in seeds, while other molecular species may dominatesignaling functions in certain tissues [19]. Indeed, the observation that NAE 12 :0 (andNAE 1 :2) have potent growth-retarding effects, when applied to young Arabidopsisseedlings (Fig. 4,a), support the contention that NAEs act as lipid mediators and haveto be depleted for normal cell expansion and seedling establishment to proceed.The effects of elevated NAEs on specific cellular responses are beginning to be

described [82] [87]. They include interference with endomembrane dynamics, alter-ations in cell shape, improper cell-wall formation, and changes in cytoskeletal structure,all of which are sure to contribute to the overall reduction in organ size and seedlinggrowth mediated by NAEs. Future work should be aimed at defining the precisemechanisms by which NAE metabolism and NAE types participate in the negativeregulation of these cellular processes and overall seedling growth. Clearly, theavailability of Arabidopsis mutant lines with altered sensitivity toward NAEs willfacilitate these studies.4.2. The Influence of NAE on Seedling Growth may be Facilitated by its Effects on

Phospholipase D and through a Remodeling of the Cytoskeleton. Interestingly, adetailed cellular analysis has shown that both the microtubule and actin cytoskeleton inArabidopsis seedlings displayed enhanced bundling and disrupted organization afterlong-term exposure to NAE 12 :0 (Fig. 4,b) [82] [87]. Such alterations in cytoskeletalorganization by NAE would likely disturb normal vesicle trafficking or cell-wallformation [88] [89], resulting in reduced cellular expansion and, consequently, indisrupted seedling development (Fig. 4,a and 4,b). This notion is further supported bythe converse observation, where overexpressing AtFAAH led to a significant increasein average cell size of seedlings, as well as a shift in the overall size distribution of cells(cotyledon epidermal cells) [55]. Although the impact of NAE on the cytoskeleton wasquite dramatic [82], it remains unclear whether such effects are the cause orconsequence of altered cellular growth.4.2.1. NAE and Microtubule Organization. Recent studies on molecules that

impinge on the plant cytoskeleton are beginning to point to potential mechanisms intohow NAEs might influence cytoskeletal structure and seedling growth. One possibilitycould be through its effect on PLD activity. As discussed in the preceding section, NAEis a potent inhibitor of PLD-a activity in vitro [80]. This finding is significant in view ofa report showing that another isoform of PLD, namely PLD-d, binds to microtubules intobacco cells [90]. Furthermore, phosphatidic acid (PA), which is generated by PLDhydrolysis of membrane phospholipids, binds to tubulin, the major protein componentof microtubules [91]. Circumstantial evidence suggests that the NAE-induced growthinhibition could, in part, be due to cytoskeletal disruption resulting from modulation ofPLD action. For example, plant cells that have been treated with butan-1-ol (BuOH) –a non-selective inhibitor of PLD activity, blocking the transphosphatidylation reaction


catalyzed by all isoforms of PLD, resulting in reduced PA production [92] – exhibiteddrastic changes in microtubule organization and inhibition of seedling growth inArabidopsis (Fig. 4,b and 4,c) [82] [93] [94]. Since the inhibition of seedling growthresulting from BuOH and NAE treatment was somewhat similar, NAE could exert itseffects on the microtubule cytoskeleton and cell expansion via inhibition of PLDactivity [87]. However, distinct differences between the effects of each compound, withNAE causing a re-organization of microtubules and BuOH-inducing depolymerizationof microtubule, suggest that NAEs have other cellular targets that are independent ofits effects on PLD activity [82]. Although there appear to be other parallels in regard tohow BuOH and NAE affect microtubule behavior (e.g., during root protoplast-volumeregulation [95]), teasing apart the precise pathways that are due to NAENs impact onPLD from other potential cellular targets, such as the NAE-binding protein [79],constitute a major challenge for the future.4.2.2. NAEs and Actin Organization. Lipid signaling in plants in part involves

interaction of PLD and phospholipid intermediates with the actin cytoskeleton viaactin-binding proteins [96]. This is most apparent in tip-growing plant cells such as roothairs, which depend on a dynamic actin network to sustain growth [97]. For example,antisense suppression of PLD-zeta-1 (PLC-z-1) [98] and a knockout to a gene involvedin phosphocholine biosynthesis [99] were shown to affect root-hair patterning inArabidopsis. Likewise, BuOH and NAE 12 :0 both inhibited root-hair elongation inArabidopsis seedlings [82] [94], and caused an apparent increase in bundling of actinfilaments along the length of the root hair [82]. These findings are similar to BuOH-induced growth inhibition and actin bundling at the pollen apex [100]. Although, theprecise mechanisms in which BuOH and NAE influence actin dynamics are not known,PA-induced inhibition of heterotrimeric actin-capping protein from Arabidopsis [101]suggest a role for lipid signaling in modulating plant cytoskeletal organization. Futurestudies are needed to determine whether the effect of NAE on actin organization isfacilitated by its impact on actin-regulatory proteins, since several of these proteins caninteract with lipids, and PLD itself has been shown to associate with actin[96] [102] [103].4.3.NAEsMay Regulate Seedling Growth and Response to Abiotic Stress through Its

Interaction with Other Plant-Signaling Pathways. The observed effects of NAE oncytoskeletal organization and PLD, both of which are known to affect a wide range ofbiological processes in plants [104] [105], have opened the possibility that, like inmammalian systems, NAEs could impinge upon a variety of plant-signaling pathways toexert its biological effects. Coincidently, many of the molecular targets and componentsof endocannabinoid signaling in animals – including ion channels, nitric oxide (NO),reactive oxygen species (ROS), G-proteins, and glutamate receptors [106] – have nowbeen implicated in regulating important plant-physiological processes. Below, we willsuggest the possibility that NAE might use some of the same targets of mammalianendocannabinoid signaling in regulating plant development and stress responses[22] [97] [107].4.3.1. Glutamate Signaling and NAE Metabolism in Plants. As already noted by

several articles in this volume, endocannabinoids exert their functions in the brain viaretrograde signaling, where postsynaptic depolarization triggers the opening of voltage-gated Ca2þ channels. An increase in cytosolic Ca2þ then activates enzymes that


synthesize anandamide. The synthesis of endocannabinoids also can be triggered bymetabotropic or ionotropic glutamate receptors that facilitate the entry of Ca2þ intothe cell [108] [109]. In the last few years, the discovery that the Arabidopsis genomeencodes ca. 20 plant ionotropic glutamate receptors [110] initiated a flurry of researchto probe deeper into the functional role for this group of plant proteins. One of the firstindications that glutamate might be exerting some sort of physiological effects in plantcells, that parallel its chemical messenger function in animals, came from observationsthat transient changes in cytosolic Ca2þ in plants are induced by glutamate application[111] [112]. The recent finding that the Arabidopsis homolog of an animal ionotropicglutamate receptor (GLR3.3) was shown to modulate Ca2þ changes and membranedepolarization in roots now provide a direct link between ion-channel activity andglutamate receptors [113].Interestingly, some of the effects of NAE on seedling development appear to be

impacted in a similar manner by glutamate application. For instance, when l-glutamatewas applied to Arabidopsis seedlings, primary root growth was inhibited, and lateralroot formation was promoted [114] in a manner reminiscent of exogenous applicationof NAE 12 :0 [87], or by exposure to the structurally related alkamides [107] [115].Furthermore, a T-DNA insertion mutant to a glutamate-receptor-like protein in rice(GLR3.1) resulted in reduced root elongation, which was due to a defect in mitoticactivity in the root apex [116]. Interestingly, glutamate was shown to depolymerizemicrotubules in roots of Arabidopsis, and this effect was blocked by antagonists ofmammalian ionotropic glutamate receptors [117], raising the possibility that glutamatesignaling in plants, like NAE action, involves modifications in cytoskeletal function.Although the parallels between exogenous glutamate and NAE on root growth arequite striking, we know nothing about how glutamate signaling is linked to the observedbiological effects of NAEs. In mammalian systems, anandamide, synthesized by thepost-synaptic neuron in response to elevated Ca2þ levels, binds to G-protein-coupledreceptors in the pre-synaptic cell, which then relays information to the pre-synaptic cellto stop releasing glutamate [109]. To establish a link between NAE and glutamatesignaling in plants, it would be beneficial to determine how glutamate applicationaffects NAE synthesis, and whether NAE influences the glutamate-induced Ca2þ

transients in roots [113].4.3.2. NAE Modulation of Abiotic-Stress Responses via PLD and Abscisic Acid

(ABA). The fact that NAE is linked to PLD as an inhibitor [80], as well as a product ofPLD hydrolytic activity [46], raises the possibility that NAEs might facilitate certainaspects of abiotic-stress responses of plants, for which PLD has already beenimplicated, including responses to cold, drought, wounding, and pathogens[104] [105]. Recent studies are beginning to point to similar roles for NAEs inmediating both biotic- (see below) and abiotic-stress responses. In regard to abioticstress, NAPE, which is the immediate precursor of NAE, was shown to accumulateduring anoxic stress in potato cells [9]. Moreover, there is recent evidence showing thatNAE delays the senescence of cut flowers by up-regulating the activity of enzymesinvolved in antioxidant defense [118]. Thus, these studies indicate that, like inmammalian systems, the build up of NAEs during abiotic stress could be requiredfor the maintenance of membrane integrity when the cell is subjected to stress[16] [22].


It is also well-established that many of the abiotic-stress responses that areregulated by PLD involve the plant hormone ABA [104] [105]. Interestingly, ABAsignaling in plants shares common features with NAE signaling in mammals, includingthe involvement of the glutamate receptors discussed above [119] as well assphingolipids [120] [121]. Also worth noting is the fact that ABA modulates seedgermination through heterotrimeric G-proteins [122] [123], while binding of NAEs toG-protein-coupled receptors in the mammalian central nervous system (CNS) is onemechanism by which NAE exerts its physiological roles [106] [109]. While we can onlyspeculate about the functions of NAEs in plant abiotic-stress responses, the observationthat NAEs are abundant in seeds and depleted in a fashion similar to ABA suggeststhat these two metabolites might interact in seed development, germination, and earlyseedling establishment [7] [20] [124]. Also, NAE has been shown to block ABA-induced stomatal closure, which is a process facilitated by G-proteins and PLD[80] [84] [125]. Although there is reason to anticipate, based on their similar moleculartargets, that ABA and NAE do interact in regulating plant responses to abiotic stress,additional experiments are needed to unravel the precise mechanisms by which theseinteractions occur.4.4. Function of NAE Metabolism in Plant–Pathogen Interactions. In animal

systems, NAEs are up-regulated under a variety of pathophysiological conditions,including immunological stresses [8] [21] [126] [127], liver diseases [128], gastrointes-tinal inflammatory disorders [129], and cardiovascular activities [130] [131]. In plants,NAEs accumulate in damaged tissues and may contribute to cytoprotection as well asin the activation of plant-defense genes [10] [18] [27]. Here, we will specifically discussthe potential role of NAEs in signal perception in response to plant pathogens.Studies with tobacco-cell suspensions and xylanase, an active molecular component

of fungal pathogens, revealed a marked change in NAE metabolism [10] [27]. Thexylanase–tobacco model system has been used extensively to study signal-transductionevents involving pathogen perception, since certain tobacco cultivars express a receptorfor xylanase, which, upon recognition of xylanase, elicits an array of responsescharacteristic of plant defense [132] [133]. In response to xylanase treatment, there wasan extracellular accumulation of NAE 14 :0 and 12 :0 in tobacco-cell-suspensioncultures [10] [27]. In additional studies with whole plants, the levels of NAE 14 :0increased 10- to 50-fold (from 6.0�4 to 64�29 ng/g fresh weight) in leaves infiltratedfor 10 min with xylanase [18], which implies a role for NAEs in defense-signaltransduction. Further support for NAE involvement in response to biotic stress wasbased on two well-documented defense responses, the induction of ion fluxes [134] andactivation of defense-gene expression [135]. Exogenously applied NAEs (NAE 14 :0,among other compounds) inhibited the alkalinization response [136] induced by avariety of fungal and bacterial elicitors (xylanase, cryptogein, harpin, and ergosterol) ina time- and concentration-dependent manner, and also activated the expression ofPAL2, a defense gene expressed in tobacco in response to xylanase (and pathogens ingeneral). Collectively, these results indicate NAE metabolism in a signal-transductionpathway that modulates cellular-defense responses following the perception ofpathogens. Furthermore, Arabidopsis plants with altered AtFAAH expression had lessamounts of total NAEs, when compared to wild-type plants [20], and these plants werehyper-susceptible to both host and non-host pathogens (L. Kang et al., work in


preparation). These data further support a role for NAEs in plant immunity, andprovide a plant system with perturbed NAE metabolism, wherein precise mechanismscan be identified.

5. Significance of NAEMetabolism in Plants. – NAEs occur in different amounts atdifferent stages of plant development, and NAE metabolism is mostly conservedbetween plants and animals. Application of NAEs to plant tissues generally leads toreduced growth and/or increased stress responses depending upon the NAE type andplant tissue/developmental stage, suggesting that the metabolism of NAEs is animportant regulatory pathway in plants. Recent experiments in Arabidopsis lines withaltered FAAH expression corroborate conclusions from pharmacological approaches,and support the concept that plant growth is negatively regulated by NAEs. In addition,evidence is accumulating, particularly in FAAH overexpressors, that the perturbationof endogenous NAE metabolism leads to enhanced growth and suppression ofresponses by plants to abiotic and biotic stressors, indicating that NAE metabolismresides at the balance between plant growth and defense (Fig. 5). While many cellulareffects have been described for NAEs, the primary targets of NAE perception/action invivo remain to be elucidated, and a clear relationship between changes in endogenousNAE profiles and physiological responses is still lacking. Although much remains to bedone, collectively the current evidence firmly places NAE lipid mediators as importantregulators in plant physiology, and their metabolism is part of an NAE regulatorypathway with striking similarities to the endocannabinoid regulatory pathway invertebrates.

Fig. 5. Summary diagram for the hypothetical placement of N-acylethanolamine (NAE) metabolism inplants at the balance between growth cues and defense cues. Decreased FAAH activity and turnover ofNAEs (or exogenous application of NAEs) leads to an increase in steady-state NAE levels, whichpromotes reduced growth, but enhances defense responses. Conversely, increased FAAH activity andturnover of NAEs (like with FAAH overexpression) leads to a reduction in steady-state NAE levels,which promotes enhanced growth, but reduces defense responses. This model, although somewhatspeculative, is consistent with available data and provides guidance for future experimental examination

and refinement of the important functional role of NAE metabolism in plants.


We thank Dr. Charlene Case for help in preparing the manuscript and figures. Work on NAEmetabolism in the laboratories ofE. B. B. andK. D. C. is supported by a grant from theUSDepartment ofEnergy, section Energy Biosciences (DE-FG02-05ER15647).

REFERENCES

[1] H. H. Schmid, P. C. Schmid, V. Natarajan, Prog. Lipid Res. 1990, 29, 1.[2] K. D. Chapman, T. S. Moore Jr., Plant Physiol. 1993, 102, 761.[3] K. D. Chapman, I. Lin, A. D. DeSouza, Arch. Biochem. Biophys. 1995, 318, 401.[4] K. D. Chapman, W. B. Sprinkle, Plant Physiol. 1995, 108, 128.[5] K. D. Chapman, W. B. Sprinkle, J. Plant Physiol. 1996, 149, 277.[6] J. A. Sandoval, Z. H. Huang, D. C. Garrett, D. A. Gage, K. D. Chapman, Plant Physiol. 1995, 109,

269.[7] K. D. Chapman, B. J. Venables, R. Markovic, R. W. Blair Jr., C. Bettinger, Plant Physiol. 1999, 120,

1157.[8] H. H. Schmid, P. C. Schmid, V. Natarajan, Chem. Phys. Lipids 1996, 80, 133.[9] A. J. Rawyler, R. A. Braendle, Plant Physiol. 2001, 127, 240.[10] K. D. Chapman, A. C. Jackson, R. A. Moreau, S. Tripathy, Physiol. Plant. 1995, 95, 120.[11] F. A. Kuehl, T. A. Jacob, O. H. Ganley, R. E. Ormond, M. A. P. Meisenger, J. Am. Chem. Soc. 1957,

79, 5577.[12] W. A. Devane, L. Hanus, A. Breuer, R. G. Pertwee, L. A. Stevenson, G. Griffin, D. Gibson, A.

Mandelbaum, A. Etinger, R. Mechoulam, Science 1992, 258, 1946.[13] A. Giuffrida, D. Piomelli, FEBS Lett. 1998, 422, 373.[14] A. Giuffrida, F. Rodriguez de Fonseca, D. Piomelli, Anal. Biochem. 2000, 280, 87.[15] P. C. Schmid, K. D. Schwartz, C. N. Smith, R. J. Krebsbach, E. V. Berdyshev, H. H. Schmid, Chem.

Phys. Lipids 2000, 104, 185.[16] K. D. Chapman, Chem. Phys. Lipids 2000, 108, 221.[17] K. D. Chapman, B. J. Venables, E. E. Dian, G. W. Grossa, J. Am. Oil Chem. Soc. 2003, 80, 223.[18] S. Tripathy, B. J. Venables, K. D. Chapman, Plant Physiol. 1999, 121, 1299.[19] B. J. Venables, C. A. Waggoner, K. D. Chapman, Phytochemistry 2005, 66, 1913.[20] Y. S. Wang, R. Shrestha, A. Kilaru, W. Wiant, B. J. Venables, K. D. Chapman, E. B. Blancaflor, Proc.

Natl. Acad. Sci. U.S.A. 2006, 103, 12197.[21] V. Di Marzo, Biochim. Biophys. Acta 1998, 1392, 153.[22] K. D. Chapman, Prog. Lipid Res. 2004, 43, 302.[23] E. V. Berdyshev, Chem. Phys. Lipids 2000, 108, 169.[24] A. F. Coburn, C. E. Graham, J. Haninger, J. Exp. Med. 1954, 100, 425.[25] J. L. Verme, J. Fu, G. Astarita, G. La Rana, R. Russo, A. Calignano, D. Piomelli, Mol. Pharmacol.

2005, 67, 15.[26] B. F. Cravatt, A. H. Lichtman, Curr. Opin. Chem. Biol. 2003, 7, 469.[27] K. D. Chapman, S. Tripathy, B. Venables, A. D. Desouza, Plant Physiol. 1998, 116, 1163.[28] H. H. O. Schmid, P. C. Schmid, E. V. Berdyshev, Chem. Phys. Lipids 2002, 121, 111.[29] B. F. Cravatt, D. K. Giang, S. P. Mayfield, D. L. Boger, R. A. Lerner, N. B. Gilula, Nature 1996, 384,

83.[30] M. K. McKinney, B. F. Cravatt, Annu. Rev. Biochem. 2005, 74, 411.[31] R. Shrestha, R. A. Dixon, K. D. Chapman, J. Biol. Chem. 2003, 278, 34990.[32] B. Q. Wei, T. S. Mikkelsen, M. K. McKinney, E. S. Lander, B. F. Cravatt, J. Biol. Chem. 2006, 281,

36569.[33] R. Shrestha, M. A. Noordermeer, M. van der Stelt, G. A. Veldink, K. D. Chapman, Plant Physiol.

2002, 130, 391.[34] M. van der Stelt, M. A. Noordermeer, T. Kiss, G. van Zadelhoff, B. Merghart, G. A. Veldink, J. F. G.

Vliegenthart, Eur. J. Biochem. 2000, 267, 2000.[35] K. D. Chapman, T. S. Moore Jr., Arch. Biochem. Biophys. 1993, 301, 21.


[36] K. D. Chapman, T. S. Moore Jr., Biochim. Biophys. Acta 1994, 1211, 29.[37] H. H. O. Schmid, E. V. Berdyshev, Prostaglandins Leukot. Essent. Fatty Acids 2002, 66, 363.[38] Y. Okamoto, J. Morishita, K. Tsuboi, T. Tonai, N. Ueda, J. Biol. Chem. 2004, 279, 5298.[39] Y. Okamoto, J. Morishita, J. Wang, P. C. Schmidt, R. J. Krebsbach, H. H. O. Schmidt, N. Ueda,

Biochem. J. 2005, 389, 241.[40] G. Petersen, H. S. Hansen, FEBS Lett. 1999, 455, 41.[41] P. C. Schmid, P. V. Reddy, V. Natarajan, H. H. Schmid, J. Biol. Chem. 1983, 258, 9302.[42] J. Morishita, Y. Okamoto, K. Tsuboi, M. Ueno, H. Sakamoto, N. Maekawa, N. Ueda, J. Neurochem.

2005, 94, 753.[43] O. Merkel, P. C. Schmid, F. Paltauf, H. H. O. Schmid, Biochim. Biophys. Acta: Mol. Cell Biol. Lipids

2005, 1734, 215.[44] E. B. Blancaflor, K. D. Chapman, in MCommunication in Plants: Neuronal Aspects of Plant LifeN,

Eds. F. Baluska, S. Mancuso, D. Volkmann, Spinger-Verlag, Heidelberg, 2006, pp. 205–219.[45] X. Wang, Prog. Lipid Res. 2000, 39, 109.[46] K. Pappan, S. Austin-Brown, K. D. Chapman, X. Wang, Arch. Biochem. Biophys. 1998, 353, 131.[47] G. Petersen, K. D. Chapman, H. S. Hansen, J. Lipid Res. 2000, 41, 1532.[48] D. Leung, A. Saghatelian, G. M. Simon, B. F. Cravatt, Biochemistry 2006, 45, 4720.[49] G. M. Simon, B. F. Cravatt, J. Biol. Chem. 2006, 281, 26465.[50] J. Liu, L.Wang, J. Harvey-White, D. Osei-Hyiaman, R. Razdan, Q. Gong, A. C. Chan, Z. Zhou, B. X.

Huang, H. Y. Kim, G. Kunos, Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 13345.[51] M. H. Bracey, M. A. Hanson, K. R. Masuda, R. C. Stevens, B. F. Cravatt, Science 2002, 298, 1793.[52] R. Shrestha, S. C. Kim, J. M. Dyer, R. A. Dixon, K. D. Chapman, Biochim. Biophys. Acta: Mol. Cell

Biol. Lipids 2006, 1761, 324.[53] N. Ueda, Prostaglandins Other Lipid Mediat. 2002, 68–69, 521.[54] N. Ueda, R. A. Puffenbarger, S. Yamamoto, D. G. Deutsch, Chem. Phys. Lipids 2000, 108, 107.[55] J. Wang, Y. Okamoto, J. Morishita, K. Tsuboi, A. Miyatake, N. Ueda, J. Biol. Chem. 2006, 281, 12325.[56] B. F. Cravatt, K. Demarest, M. P. Patricelli, M. H. Bracey, D. K. Giang, B. R. Martin, A. H.

Lichtman, Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 9371.[57] Y. X. Sun, K. U. Tsuboi, L. Y. Zhao, Y. Okamoto, D. M. Lambert, N. Ueda, Biochim. Biophys. Acta:

Mol. Cell Biol. Lipids 2005, 1736, 211.[58] K. Tsuboi, Y. X. Sun, Y. Okamoto, N. Araki, T. Tonai, N. Ueda, J. Biol. Chem. 2005, 280, 11082.[59] A. M. Mulder, B. F. Cravatt, Biochemistry 2006, 45, 11267.[60] T. M. Dunn, D. V. Lynch, L. V. Michaelson, J. A. Napier, Ann. Bot. (London) 2004, 93, 483.[61] S. Pollmann, D. Neu, T. Lehmann, O. Berkowitz, T. Schafer, E. W. Weiler, Planta 2006, 224, 1241.[62] R. D. Page, Comput. Appl. Biosci. 1996, 12, 357.[63] E. Blee, Prog. Lipid Res. 1998, 37, 33.[64] A. Liavonchanka, I. Feussner, J. Plant Physiol. 2006, 163, 348.[65] G. van Zadelhoff, G. A. Veldink, J. F. Vliegenthart, Biochem. Biophys. Res. Commun. 1998, 248, 33.[66] M. van der Stelt, W. F. Nieuwenhuizen, G. A. Veldink, J. F. G. Vliegenthart, FEBS Lett. 1997, 411,

287.[67] N. Ueda, Y. Kurahashi, S. Yamamoto, T. Tokunaga, J. Biol. Chem. 1995, 270, 23823.[68] K. R. Kozak, L. J. Marnett, Prostaglandins Leukot. Essent. Fatty Acids 2002, 66, 211.[69] R. G. Pertwee, Int. J. Obesity (London) 2006, 30 (Suppl. 1) , S13.[70] T. Sugiura, S. Kishimoto, S. Oka, M. Gokoh, Prog. Lipid Res. 2006, 45, 405.[71] P. H. Reggio, Handb. Exp. Pharmacol. 2005, 247.[72] S. Tripathy, K. Kleppinger-Sparace, R. A. Dixon, K. D. Chapman, Plant Physiol. 2003, 131, 1781.[73] C. J. Hillard, W. S. Edgemond, W. B. Campbell, J. Neurochem. 1995, 64, 677.[74] S. Bourqe, M. N. Binet, M. Ponchet, A. Pugin, A. Lebrun-Garcia, J. Biol. Chem. 1999, 274, 34699.[75] S. Bourqe, M. Ponchet, M. N. Binet, P. Ricci, A. Pugin, A. Lebrun-Garcia, Plant Physiol. 1998, 118,

1317.[76] J. Lee, D. F. Klessig, T. Nurnberger, Plant Cell 2001, 13, 1079.[77] A. D. Khanolkar, S. L. Palmer, A. Makriyannis, Chem. Phys. Lipids 2000, 108, 37.[78] T. Sugiura, Y. Kobayashi, S. Oka, K. Waku, Prostaglandins Leukot. Essent. Fatty Acids 2002, 66, 173.


[79] S. Tripathy, K. Kleppinger-Sparace, R. A. Dixon, K. D. Chapman, in MAdvanced Research on PlantLipidsN, Eds. N. Murata, M. Yamada, I. Nishida, H. Okumyama, J. Sekiya, W. Hajime, KluwerAcademic Publishers, Dordrecht, 2003, p. 315–318.

[80] S. L. Austin-Brown, K. D. Chapman, Plant Physiol. 2002, 129, 1892.[81] L. Fan, S. Zheng, X. Wang, Plant Cell 1997, 9, 2183.[82] C. M. Motes, P. Pechter, C. M. Yoo, Y. S. Wang, K. D. Chapman, E. B. Blancaflor, Protoplasma 2005,

226, 109.[83] A. M. Jones, S. M. Assmann, EMBO Rep. 2004, 5, 572.[84] G. Mishra, W. Zhang, F. Deng, J. Zhao, X. Wang, Science 2006, 312, 264.[85] S. Ritchie, S. Gilroy, Plant Physiol. 2000, 124, 693.[86] S. V. Siegmund, E. Seki, Y. Osawa, H. Uchinami, B. F. Cravatt, R. F. Schwabe, J. Biol. Chem. 2006,

281, 10431.[87] E. B. Blancaflor, G. Hou, K. D. Chapman, Planta (Berlin) 2003, 217, 206.[88] A. R. Paredez, C. R. Somerville, D. W. Ehrhardt, Science 2006, 312, 1491.[89] B. Voigt, A. C. Timmers, J. Samaj, A. Hlavacka, T. Ueda, M. Preuss, E. Nielsen, J. Mathur, N. Emans,

H. Stenmark, A. Nakano, F. Baluska, D. Menzel, Eur. J. Cell Biol. 2005, 84, 609.[90] J. C. Gardiner, J. D. Harper, N. D.Weerakoon, D. A. Collings, S. Ritchie, S. Gilroy, R. J. Cyr, J. Marc,

Plant Cell 2001, 13, 2143.[91] C. Testerink, H. L. Dekker, Z. Y. Lim, M. K. Johns, A. B. Holmes, C. G. Koster, N. T. Ktistakis, T.

Munnik, Plant J. 2004, 39, 527.[92] T. Jacob, S. Ritchie, S. M. Assmann, S. Gilroy, Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 12192.[93] P. Dhonukshe, A. M. Laxalt, J. Goedhart, T. W. Gadella, T. Munnik, Plant Cell 2003, 15, 2666.[94] J. Gardiner, D. A. Collings, J. D. Harper, J. Marc, Plant Cell Physiol. 2003, 44, 687.[95] G. Komis, H. Quader, B. Galatis, P. Apostolakos, New Phytol. 2006, 171, 737.[96] B. K. Drobak, V. E. Franklin-Tong, C. J. Staiger, New Phytol. 2004, 163, 13.[97] E. B. Blancaflor, Y. S. Wang, C. M. Motes, Int. Rev. Cytol. 2006, 252, 219.[98] Y. Ohashi, A. Oka, R. Rodrigues-Pousada, M. Possenti, I. Ruberti, G. Morelli, T. Aoyama, Science

2003, 300, 1427.[99] A. Cruz-Ramirez, J. Lopez-Bucio, G. Ramirez-Pimentel, A. Zurita-Silva, L. Sanchez-Calderon, E.

Ramirez-Chavez, E. Gonzalez-Ortega, L. Herrera-Estrella, Plant Cell 2004, 16, 2020.[100] D. Monteiro, Q. Liu, S. Lisboa, G. E. Scherer, H. Quader, R. Malho, J. Exp. Bot. 2005, 56, 1665.[101] S. Huang, L. Gao, L. Blanchoin, C. J. Staiger, Mol. Biol. Cell 2006, 17, 1946.[102] D. J. Kusner, J. A. Barton, C. Qin, X. Wang, S. S. Iyer, Arch. Biochem. Biophys. 2003, 412, 231.[103] D. J. Kusner, J. A. Barton, K. K. Wen, X. Wang, P. A. Rubenstein, S. S. Iyer, J. Biol. Chem. 2002, 277,

50683.[104] B. O. Bargmann, T. Munnik, Curr. Opin. Plant Biol. 2006, 9, 515.[105] X. Wang, Plant Physiol. 2005, 139, 566.[106] C. J. Fowler, Brain Res. Brain Res. Rev. 2003, 41, 26.[107] J. Lopez-Bucio, G. Acevedo-Hernandez, E. Ramirez-Chavez, J. Molina-Torres, L. Herrera-Estrella,

Curr. Opin. Plant Biol. 2006, 9, 523.[108] S. P. Brown, S. D. Brenowitz, W. G. Regehr, Nat. Neurosci. 2003, 6, 1048.[109] R. I. Wilson, R. A. Nicoll, Science 2002, 296, 678.[110] H. M. Lam, J. Chiu, M. H. Hsieh, L. Meisel, I. C. Oliveira, M. Shin, G. Coruzzi, Nature 1998, 396,

125.[111] K. L. Dennison, E. P. Spalding, Plant Physiol. 2000, 124, 1511.[112] O. Meyerhoff, K. Muller, M. R. Roelfsema, A. Latz, B. Lacombe, R. Hedrich, P. Dietrich, D. Becker,

Planta 2005, 222, 418.[113] Z. Qi, N. R. Stephens, E. P. Spalding, Plant Physiol. 2006, 142, 963.[114] P. Walch-Liu, L. H. Liu, T. Remans, M. Tester, B. G. Forde, Plant Cell Physiol. 2006, 47, 1045.[115] E. Ramirez-Chavez, J. Lopez-Bucio, L. Herrera-Estrella, J. Molina-Torres, Plant Physiol. 2004, 134,

1058.[116] J. Li, S. Zhu, X. Song, Y. Shen, H. Chen, J. Yu, K. Yi, Y. Liu, V. J. Karplus, P. Wu, X. W. Deng, Plant

Cell 2006, 18, 340.


[117] M. Sivaguru, S. Pike, W. Gassmann, T. I. Baskin, Plant Cell Physiol. 2003, 44, 667.[118] Y. Zhang, W.-M. Guo, S.-M. Chen, L. Han, Z.-M. Li, J. Plant Physiol. 2007, in press (doi: 10.1016/

j.jplph.2006.07.003).[119] J. Kang, S. Mehta, F. J. Turano, Plant Cell Physiol. 2004, 45, 1380.[120] S. Coursol, L. M. Fan, H. Le Stunff, S. Spiegel, S. Gilroy, S. M. Assmann, Nature 2003, 423, 651.[121] S. Coursol, H. Le Stunff, D. V. Lynch, S. Gilroy, S. M. Assmann, S. Spiegel, Plant Physiol. 2005, 137,

724.[122] S. Pandey, S. M. Assmann, Plant Cell 2004, 16, 1616.[123] S. Pandey, J. G. Chen, A. M. Jones, S. M. Assmann, Plant Physiol. 2006, 141, 243.[124] J. V. Jacobsen, D. W. Pearce, A. T. Poole, R. P. Pharis, L. N. Mander, Physiol. Plant 2002, 115, 428.[125] W. Zhang, C. Qin, J. Zhao, X. Wang, Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 9508.[126] E. J. Carrier, J. A. Auchampach, C. J. Hillard, Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 7895.[127] J. Fernandez-Ruiz, F. Berrendero, M. L. Hernandez, J. A. Ramos, Trends Neurosci. 2000, 23, 14.[128] A. Mallat, S. Lotersztajn, J. Endocrinol. Invest. 2006, 29, 58.[129] F. Massa, M. Storr, B. Lutz, J. Mol. Med. 2005, 83, 944.[130] C. R. Hiley, W. R. Ford, Biol. Rev. Camb. Philos. Soc. 2004, 79, 187.[131] P. Pacher, S. Batkai, G. Kunos, Handb. Exp. Pharmacol. 2005, 599.[132] U. Hanania, A. Avni, Plant J. 1997, 12, 113.[133] N. Matarasso, S. Schuster, A. Avni, Plant Cell 2005, 17, 1205.[134] J. Ebel, D. Scheel, MElicitor Recognition and Signal TransductionN, in MGenes Involved in Plant

DefenseN, Eds. T. Boller, F. Meins, Springer-Verlag, New York, 1992, pp. 183–205.[135] R. A. Dixon, C. J. Lamb, Annu. Rev. Plant Physiol. Plant Mol. Biol. 1990, 41, 339.[136] N. Furman-Matarasso, E. Cohen, Q. Du, N. Chejanovsky, U. Hanania, A. Avni, Plant Physiol. 1999,

121, 345.

Received December 9, 2006


Documents

The N-Acylethanolamine-Mediated Regulatory Pathway in Plants