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Analytical and Bioanalytical Chemistry

, Volume 408, Issue 22, pp 6141–6151 | Cite as

Identification of the oleic acid ethanolamide (OEA) isomer cis-vaccenic acid ethanolamide (VEA) as a highly abundant 18:1 fatty acid ethanolamide in blood plasma from rats and humans

  • Waldemar Röhrig
  • Reiner Waibel
  • Christopher Perlwitz
  • Monika Pischetsrieder
  • Tobias HochEmail author
Research Paper

Abstract

The endocannabinoid system is important in various physiological pathways, especially the regulation of food intake. It consists of endocannabinoids like 2-arachidonoyl-glycerol (2-AG) or the fatty acid ethanolamide archachidonoyl-ethanolamide (AEA) with binding affinity to cannabinoid receptors. Further, fatty acid ethanolamides (FAEAs) influence the endocannabinoid system without affecting cannabinoid receptors by using independent physiological pathways. Among FAEAs, oleic acid ethanolamide (OEA) gained importance because of its promising ability to reduce food intake. By ultrahigh-performance liquid chromatography–electrospray ionization–tandem mass spectrometry (UHPLC–ESI–MS/MS), we detected a chromatographically separated molecule in plasma samples from rats and humans with identical mass and fragmentation patterns as those of OEA. Via synthesis and extensive analysis of ethanolamides of different cis/trans- and position isomers of oleic acid (cis9-18:1), we could identify the unknown molecule as vaccenic acid (cis11-18:1) ethanolamide (VEA). In this study we identified VEA as the most abundant 18:1 FAEA in rat plasma and the second most abundant 18:1 FAEA in human plasma.

Keywords

OEA VEA Fatty acid ethanolamides Endocannabinoid system LC–MS/MS 

Introduction

The endocannabinoid system consists of endocannabinoids and their structurally related fatty acid ethanolamide (FAEA) species, the cannabinoid receptors CB1 and CB2 [1, 2], and metabolic enzymes, which are responsible for the formation and the degradation of endocannabinoids and endocannabinoid-like structures [3]. The endocannabinoids 2-arachidonoyl-glycerol (2-AG) [4], N-arachidonoyl ethanolamine (anandamide) [5], or noladin ether have been identified as naturally occurring cannabinoid receptor agonists [6], whereas the naturally occurring N-arachidonyl dopamine is a synthetic CB1 agonist [7] and O-arachidonoyl ethanolamine (virodhamine) acts both as an antagonist at CB1 and as an agonist at CB2 receptors [8].

Endocannabinoids and related compounds play decisive roles in different physiological pathways like the control of pain [9] or REM sleep [10]. Furthermore, food intake and energy balance are regulated by the endocannabinoid system [11], e.g., via enhanced olfactory perception [12]. Whereas AEA has a stimulating effect on food intake by interacting with CB receptors, the FAEA oleic acid ethanolamide (OEA) recently gained interest because of its participation in feeding regulation and the physiological system for monitoring the dietary fat intake without acting directly at cannabinoid receptors [13]. Like other FAEAs that develop from different fatty acids, OEA is synthesized in vivo from food-derived oleic acid [14] via N-acyltransferase-catalyzed formation of N-fatty acid phosphatidylethanolamine and cleavage by phospholipase D [3].

Oleic acid is a common constituent of several fats and oils like olive oil, peanut oil, rapeseed oil, lard, or sunflower oil [15]. Additionally, oleic acid is the main fatty acid in mutton, veal, beef, or pork [9, 15]. The presence in many frequently ingested foods makes it highly available for the endogenous synthesis of the respective FAEA.

When we analyzed OEA in plasma of rats and humans, a chromatographically separated substance became apparent during electrospray ionization tandem mass spectrometry (ESI–MS/MS) with the same mass and fragmentation pattern as OEA. To elucidate this hitherto undescribed compound, we searched for other monounsaturated octadecanoic fatty acids (18:1) present in food that could also form the respective FAEA and would behave similarly during ESI–MS/MS analysis. As candidates qualify the position isomers of oleic acid (cis9-18:1), namely vaccenic acid (cis11-18:1) and petroselinic acid (cis6-18:1), or the respective trans-isomers elaidic acid (trans9-18:1), trans-vaccenic acid (trans11-18:1), and trans-petroselinic acid (trans6-18:1). Vaccenic acid mainly appears in milk, e.g., from yaks [16] or milk products like Scamorza ewe milk cheese [17]. Furthermore, vaccenic acid seems to influence the endocannabinoid system by reducing the plasma level of AEA in hypercholesterolemic subjects [18]. Petroselinic acid has been isolated from plants like coriander [19] or parsley [20]. Monounsaturated fatty acids are converted to their trans-isomers, e.g., by bio-hydrogenation in ruminants, partial hydrogenation of vegetable oils, or further processing steps [21]. The contents of trans-fatty acids in meat or products from ruminants like milk, cheese, or butter is about 5–10 %, with elaidic acid and trans-vaccenic acid as the main trans-fatty acids [22]. Additionally, food processing leads to the formation of trans-fatty acids in several foods [23]. Because position isomers as well as the respective trans-isomers from oleic acid are present in food sources, they could consequently be incorporated into the endocannabinoid system. Therefore, we aimed to identify the newly found structure by synthesizing the respective ethanolamides derived from oleic acid (OEA) and its isomers vaccenic acid (VEA), petroselinic acid (PeEA), elaidic acid (EEA), trans-vaccenic acid (tVEA), and trans-petroselinic acid (tPeEA). The structures of these ethanolamides are summarized in Fig. 1.
Fig. 1

Chemical structures of the 18:1 fatty acid ethanolamides oleic acid ethanolamide (a, OEA, cis9-18:1), elaidic acid ethanolamide (b, EEA, trans9-18:1), vaccenic acid ethanolamide (c, VEA, cis11-18:1), trans-vaccenic acid ethanolamide (d, tVEA, trans11-18:1), petroselinic acid ethanolamide (e, PeEA, cis6-18:1), and trans-petroselinic acid ethanolamide (f, tPeEA, trans6-18:1)

Recently, methods for the quantification of endocannabinoids and endocannabinoid-related compounds like FAEA including OEA have been developed to simultaneously quantify endocannabinoids and related compounds in milk and other biofluids [24], in human blood cells and human blood plasma [25, 26], or in other human bio-matrices [27] by ultrahigh-performance liquid chromatography (UHPLC)–ESI–MS/MS. Since structural differences in FAEAs can lead to different physiological effects, we consider it important to analyze OEA and its C18:1 position- and configuration isomers by comparing the respective pure compounds with those found in blood plasma from rats and humans using UHPLC–ESI–MS/MS. To the best of our knowledge, no endocannabinoids other than OEA that are based on monounsaturated octadecanoate backbones have been described to date in literature or textbooks [28].

Materials and methods

Materials

Methyl esters of fatty acids (>98 %), ethanolamine (>99 %) ethanolamine-d4 (ethanol-1,1,2,2-d4-amine, 98 atomic % D), hexamethyldisilazane (>99 %), dimethyldisulfide (>99 %), and all other reagents (of at least analytical grade) were supplied by Sigma-Aldrich Chemie GmbH (Taufkirchen, Germany). The following products were used as solvents and eluents: methanol, acetonitrile (both HiPerSolv LC–MS/MS grade, VWR International GmbH, Ismaning, Germany), formic acid (LC–MS/MS grade, Fluka, Buchs, Switzerland), toluene (Chromasolv) and dichloromethane (both Sigma-Aldrich), chloroform (analytical reagent, Fisher Scientific GmbH, Schwerte, Germany), and water from a Millipore Synergy 185 lab-water system (Merck Chemicals GmbH, Schwalbach, Germany).

Synthesis of FAEAs

The applied reaction conditions were adopted from Wang et al. [29]. Briefly, ethanolamine (1.8 mmol) was mixed with the respective fatty acid methyl ester (0.06 mmol) in a 1.5-mL Eppendorf tube. After addition of 25 μL of sodium methoxide in methanol (4.8 M), the tube was placed on a vortex mixer (VWR International) at medium speed for 30 min at room temperature. Subsequently, methanol was added to the mixture until the formed precipitate was dissolved. After further shaking for 30 min, the added methanol was removed in vacuum, the remaining mixture diluted with chloroform to a final volume of 600 μL, mixed with 400 μL of ice-cold 0.5 M HCl, and vortexed thoroughly. The phases were separated by centrifugation at 13,000×g for 1 min and 4 °C and the process was repeated three times after discarding the aqueous layer. To remove HCl, the organic phase was washed with 5 % Na2CO3. After drying over Na2SO4, the solvent was removed in a SpeedVac concentrator (Savant SPD121P, Thermo Scientific, Dreieich, Germany) yielding 86–98 % of the desired FAEA. OEA-d4 was synthesized accordingly with oleic acid methyl ester and deuterated ethanolamine as reaction components.

Gas chromatography electron ionization mass spectrometry (GC–EI–MS)

GC–EI–MS analysis was performed on an Agilent MSD quadrupole system (GC 7890A and MSD 5975C, Agilent Technologies, Waldbronn, Germany) equipped with a Gerstel CIS 4 injection system and Gerstel MPS 2 autosampler (Gerstel, Duisburg, Germany). Data were recorded and analyzed by MSD ChemStation (Agilent Technologies, Waldbronn, Germany). The system was equipped with a DB-5 column (30 m × 0.25 mm, film thickness 0.25 μm, Agilent J&W Scientific, Santa Clara, USA) and an uncoated, deactivated fused silica capillary as a precolumn (2–3 m × 0.53 mm), which was changed regularly to avoid accumulation of impurities. Helium was used as carrier gas. The total flow of the system was 1.5 mL min-1, which was transferred into the MS unit using an uncoated, deactivated fused silica capillary (0.3–0.6 m, 0.25 mm) as transfer line. GC oven starting temperature was 40 °C, held for 2 min, then the temperature was raised at a rate of 15 °C min−1 up to 200 °C, then at 20 °C min−1 to 290 °C, and held for 10 min. Injection volumes were 1.0 μL with a split ratio of 20:1 and a split flow of 20 mL min−1 with helium as carrier gas. GC–EI mass spectra were generated in full scan mode (m/z range 40 to 1000, scan rate ∼600 Da s−1) at 70 eV ionization energy.

Derivatization of FAEAs for GC–EI–MS analysis

Anhydrous newly synthesized FAEAs (0.5 g) were placed into a 2-mL Eppendorf tube each, dissolved in 100 μL of chloroform and incubated with hexamethyldisilazane (50 μL) and 3 μL of trimethylchlorosilane for 1 h under continuous shaking. The mixture was evaporated under reduced pressure, absorbed in dichloromethane, and centrifuged briefly to precipitate ammonium chloride formed during synthesis. For GC–EI–MS analysis, the organic phase was carefully transferred to a fresh tube and diluted to a final concentration of 100μg mL−1. The remaining solvent was concentrated under reduced pressure yielding a colorless and oily product.

Double-bond derivatization by dimethyl disulfide

For the confirmation of the double bond position, a method by Dunkelblum et al. was used [30]. Anhydrous silylated FAEAs (200 μg) were placed in glass vials, dissolved in 180 μL of hexane, and treated with 100 μL of dimethyl disulfide followed by 20 μL of iodine in hexane (60 mg mL−1). The vial was sealed with a screw cap and placed in an oven at 50 °C for 12 h. The mixture was diluted with hexane to a final volume of 500 μL and washed twice with 200 μL of 5 % Na2SO3 solution to remove molecular iodine. After washing with Millipore water, the organic layer was dried over Na2SO4 and yielded 64–80 % of a yellowish solid product, all of which was dissolved in dichloromethane and diluted to a final concentration of 50 μg mL−1 prior to GC–EI–MS analysis.

Qualitative analysis of FAEAs by 1H-NMR

1H-NMR analysis on a Bruker NMR spectrometer (Avance-600 UltraShield, Switzerland) operating at 600 MHz was performed to distinguish between the ester and amide formation of the reaction products described above. Ten milligrams each of the respective FAEA was dissolved in CDCl3.

(UHPLC–)ESI–MS/MS

An API 4000 LIT mass spectrometer (AB Sciex, Foster City, CA, USA) with an ESI source (Applied Biosystems, Foster City, CA, USA) was used for analysis with the following parameters (UHPLC/direct infusion): nebulizer gas (GS1) 55/12 psig, desolvation gas (GS2) 75/0 psig, temperature (TEM) 600/550 °C, ion spray voltage (IS, positive mode) 3500/5500 V. The mass spectrometer was coupled to an infusion pump for direct infusion (Harvard Apparatus 11plus, Holliston, MA, USA) or to a UHPLC system (Dionex UltiMate 3000 RS, Thermo Fisher Scientific, Idstein, Germany) consisting of a pump with degasser, autosampler (maintained at 5 °C), and a column compartment (maintained at 40 °C). As stationary phase, a Kinetex Core Shell C18 UHPLC column (Phenomenex Aschaffenburg, Germany, 1.7 μm, 150 × 2.1 mm) with a SecurityGuard ULTRA Cartridge precolumn was used. The mobile phase consisted of 0.1 % formic acid (eluent A) and acetonitrile (eluent B) with the following gradient: 0 min (55 % B), 8.0 min (80 % B), 8.1 min (100 % B), 11.9 min (100 % B), 12.0 min (55 % B), 15.0 min (55 % B), injection volume 10 μL, and flow rate 0.5 mL min−1.

Parameters for the detection of OEA, EEA, VEA, tVEA, PeEA, and tPeEA via direct infusion are as follows: precursor ion m/z 326.3, declustering potential (DP) 60 V, collision energy (CE) 25–50 V. Parameters for the detection of OEA, EEA, VEA, tVEA, PeEA, and tPeEA by UHPLC–MS/MS are as follows: precursor ion m/z 326.3, product ions m/z 62.0/95.0, DP 70/60 V, CE 34/44 V, cell exit potential (CXP) 5/4 V. Parameters for the detection of OEA-d4 by UHPLC–MS/MS are as follows: precursor ion m/z 330.3, product ion m/z 66.1, DP 70 V, CE 28 V, CXP 4 V.

Product ion spectra of VEA and OEA in concentrated plasma (5×) and in standard samples were measured via enhanced product ion scans (EPI) using the linear ion trap function with a fill time of 20 ms and a scan rate of 1000 Da s−1 with the remaining parameters as specified above. To exclude interference by carry-over between samples, water runs were performed between selected chromatographic runs, which did not show any signal at the respective MRM transitions.

Analyst software 1.5.1 was used for data acquisition and processing. Because of a partial overlap of respective cis- and trans-isomers, peak integration was carried out manually.

Blood sampling and processing

Human venous blood, collected from the forearm of 20 healthy volunteers into K+-EDTA-containing tubes (Monovette, Sarstedt, Nümbrecht, Germany), was obtained from the local blood donor bank. Since participants are generally encouraged to eat prior to blood donation, we obtained a random sample spectrum ranging from fasted to satiated individuals. Rat blood was taken from the tail vein of six male Wistar rats (Charles River, Sulzfeld, Germany), which were anesthetized by isoflurane (initially 5 % and 1.5 % maintenance, Baxter Deutschland, Unterschleißheim, Germany). The blood was drawn into K+-EDTA-containing tubes (Multivette® 600, Sarstedt) and handled on ice until processing. The rats were maintained on standard chow (Altromin 1324, Lage, Germany) under controlled conditions.

Plasma was obtained by centrifugation of whole blood at 2000×g for 10 min at 4 °C, pooled, and then aliquoted. Samples of 50 μL each were stored in 500 μL Eppendorf tubes at −80 °C until analysis. Recently, toluene proved to be a suitable solvent for the extraction of endocannabinoids [31], which suppresses the extraction of phospholipids that disturb the ionization in the mass spectrometer [26]. For extraction, each sample was diluted with Millipore water to a final volume of 200 μL and 210 μL of toluene was added. Samples were thoroughly mixed on a vortex mixer for 4 × 15 s at maximum speed and centrifuged at 14,000×g for 10 min at 30 °C. The upper organic phase was quantitatively transferred into glass vial inserts and evaporated by a SpeedVac concentrator. The remaining analytes were dissolved in a mixture of methanol/water (50:50) with 0.1 % formic acid for further analysis by UHPLC–ESI–MS/MS.

Contamination in laboratory material has been described to be a serious problem during the analysis of FAEAs [32, 33]. Mainly palmitic acid ethanolamide or OEA have been reported to be present in tubes, reagents, or consumables. Therefore, we analyzed blank solutions of the reagents and tubes under extraction or blood sampling conditions. A marginal influence on the OEA, but not the VEA signal, could be detected. In addition, the mass spectra of the other analytes were not influenced by contamination.

To assess if the signal at the retention time of VEA was caused by an artifact of OEA during sample preparation, a standard solution of OEA in MeOH/H2O + 0.1 % formic acid containing OEA-d4 as internal standard was extracted and processed as described for plasma samples. Furthermore, blood and plasma samples of rats and humans were spiked with OEA and OEA-d4 in order to evaluate if the signal for VEA was influenced by the addition of OEA before sample preparation. Each experiment was conducted in triplicate and evaluated by comparison of the relative signal areas OEA/OEA-d4 and VEA/OEA-d4 with and without OEA addition.

Results and discussion

The endocannabinoids 2-AG and AEA as well as the endocannabinoid-like FAEAs are able to influence several physiological systems such as the networks involved in regulating pain [9], REM sleep [10] and, notably, food intake and energy balance [11]. Among these FAEAs, especially OEA gained importance because of its role in regulating food intake pathways. OEA was reported to suppress food intake by different pathways [34, 35, 36] and has since been discussed as an alternative for endocannabinoid antagonists for the control of food intake [35]. The FAEAs are synthesized from food-derived fatty acids [3], the conversion of which has been described in detail for OEA in vivo [14], but not for any other C18:1 isomer.

When we analyzed OEA in plasma from rats and humans, the chromatograms showed a second, hardly separated signal of a molecule with the same mass and the same fragmentation pattern as OEA. In addition to oleic acid (cis9-18:1), other monounsaturated fatty acids with 18 carbon atoms (18:1) are present in common foods, namely petroselinic acid (cis6-18:1), vaccenic acid (cis11-18:1), elaidic acid (trans9-18:1), trans-petroselinic acid (trans6-18:1), or trans-vaccenic acid (trans11-18:1) [15, 16, 17, 18]. These fatty acids can be considered as potential sources for the formation of FAEAs in vivo and could explain our observations by UHPLC–ESI–MS/MS. To identify the unknown signal, we investigated the respective LC–MS characteristics of the synthesized ethanolamides OEA, PeEA, VEA, EEA, tPeEA, and tVEA (Fig. 1) and extracts of plasma from rats and humans.

Synthesis of OEA and OEA-isomers

For the chemical synthesis of OEA, VEA, PeEA, EEA, tVEA, and tPeEA, we developed a one-pot reaction and cleanup method. As described by Wang et al. [29], ethanolamine can be used both as solvent and reactant during reaction with methyl linoleate without further addition of other solvents. While this protocol worked well for cis6-, cis9-, and cis11-octadecanoic acid methyl esters, the synthesis of all trans-isomers varied greatly in terms of yield and purity. Primary analysis of all synthesis products by ESI–MS via direct infusion resulted in a major signal for all ethanolamides at m/z 326.3 [M + H]+ (data not shown). However, the trans9- and trans6-isomers reproducibly showed the lowest conversion rates with a considerable amount of unreacted methyl ester in the final product with a signal at m/z 297.3 [M + H]+ (data not shown). During reaction of the trans-isomers, but not the cis-isomers, a voluminous precipitate was formed that most probably consisted of a mixture of ethanolamide crystals and unreacted methyl esters, which, presumably, was the reason for the low conversion rate. Therefore, the reaction parameters had to be improved with regard to solubility of the trans-isomers. Increasing ethanolamine to 30-fold instead of the original 10-fold excess resulted in a considerable improvement of all reactions but for the trans9-isomer. Addition of methanol to the initial reaction mixture resulted in quantitative conversion of all methyl esters to ethanolamines. Judging by direct infusion/ESI–MS analysis, none of the final products contained traces of ethanolamine shown by the absence of a signal at m/z 62.1 [M + H]+, and no unreacted methyl ester with m/z of 297.3 [M + H]+ was detectable either (data not shown). With the revised reaction setup, we extended the applicability of fatty acid ethanolamide synthesis as initially published by Wang et al. [29] to trans-fatty acids isomers. Thus, we were able to reliably synthesize a spectrum of cis- and trans-octadecanoic ethanolamide isomers with quantitative conversion rates and excellent purity without complex purification steps.

Verification of purity and identity of the synthesized analytical standards via GC–MS, dimethyl disulfide derivatization, and 1H-NMR

Prior to UHPLC–ESI–MS/MS analysis, the identity and purity of the synthesized standards were further assessed by different methods. In contrast to LC–ESI–MS/MS measurements, all compounds used during synthesis are reliably detected by GC–EI–MS with little variation of ionization efficiency between molecules, so that the purity of synthesis products can be examined with more detail and reliability. To increase the volatility of our standards, a silylation reaction adapted from Wang et al. was performed [29]. Mass spectra resulting from electron ionization were highly similar between samples. The respective chromatograms exposed one main peak for each product at a retention time of about 20.7 min (see Electronic Supplementary Material (ESM) Fig. S1). Impurities were hardly detectable. The obtained results prove the high purity of the synthesized products with purities (assessed via areas of GC–EI–MS analysis of the silylated products) of >99 % for all of the synthesized standards. The purity and identity of the synthesized compounds was further confirmed by NMR measurements as described below. The position of the double bond of the synthesized products was verified by a dimethyl disulfide derivatization method starting from the silylated ethanolamides [30]. With this technique, dimethyl disulfide selectively reacts with double bonds forming derivatives with two additional methyl sulfide groups at the position of the double bonds. During GC–EI–MS analysis, these derivatives undergo specific fragmentation preferably at the derivatized positions resulting in characteristic fragments dependent on the position of the double bond. We detected the respective characteristic fragments in the synthesized FAEAs at position 9 for OEA and EEA (fragments with m/z 173 and 228, respectively, Fig. 2a), at position 11 for VEA and tVEA (fragments with m/z 145 and 256, respectively, Fig. 2b) and at position 6 for PeEA and tPeEA (fragments with m/z 215 and 186, respectively, Fig. 2c). Consequently, we could prove the position of the double bond in the synthesized analytes and exclude that isomerization of the double bond occurred during synthesis.
Fig. 2

Determination of the position of the double bond by derivatization with dimethyl disulfide and analysis by GC–EI–MS. Molecular structures of the fragments (1st column) and GC–EI–MS spectra of the respective analytes (2nd column) are shown: a oleic acid ethanolamide (OEA) and elaidic acid ethanolamide (EEA, cis/trans9-18:1); b vaccenic acid ethanolamide (VEA) and trans- (t)VEA (cis/trans11-18:1); and c petroselinic acid ethanolamide (PeEA) and trans- (t)PeEA (cis/trans6-18:1)

Virodhamine, which differs from AEA only by the linkage between the fatty acid and ethanolamine, is known to be a CB1-receptor antagonist and CB2-receptor agonist. Whereas AEA contains an amide bond, virodhamine is linked by an ester bond [8]. Both molecules have the same mass and show similar fragmentation during ESI–MS/MS analysis so that they have to be separated chromatographically [37]. During synthesis, conversion of the ethanolamide linkage to an ester linkage may occur mainly under acidic conditions, but the reaction is reversible under mildly basic conditions [38]. Therefore, it was necessary for further assessment of product identity to determine the linkage type of the synthesized FEAEs, which was achieved by 1H-NMR. Focusing on the 1H-NMR signals for the protons of the ethanolamine element allowed for the easy identification of amide- and ester linkages of the synthesized amides (1′-CH2 and 2′-CH2 position; Fig. 1) due to their well distinguishable chemical shifts [39]. The ester bond is characterized by the chemical shifts of 4.37 for 1′-CH2 and 3.26 for 2′-CH2, whereas the amide bond leads to shifts of 3.45 and 3.75, respectively. No differences are detectable in the chemical shifts of the protons of the fatty acid chain [39]. We detected all characteristic chemical shifts for an amide-linked fatty acid with ethanolamine in accordance to literature data (Table 1). Additionally, we could identify further characteristic signals for the OH hydrogen as well as the signal for the NH hydrogen in all synthesized products. The NMR spectra did not show any signals for the respective ester bonds nor for unreacted methyl ester entities of the fatty acid methyl esters (ESM Fig. S2). The presented data confirm the identity and purity of all our synthesized products.
Table 1

Chemical shifts of the 1H-NMR experiment for the synthesized oleic acid ethanolamide (OEA), elaidic acid ethanolamide (EEA), cis-vaccenic acid ethanolamide (VEA), trans-vaccenic acid ethanolamide (tVEA), cis-petroselinic acid ethanolamide (PeEA), and trans-petroselinic acid ethanolamide (tPeEA). As reference, the chemical shifts of an ester-linked and an amide-linked product of oleic acid and ethanolamine are shown as reported by Ottria et al. [39]. Characteristic shifts for the ethanolamide entities, which could be found in our analytes, are printed in bold

H

OEA ester [39]

OEA amide [39]

OEA

EEA

VEA

tVEA

PeEA

tPeEA

1′-CH 2

3.45 (dt a, 4.5)

3.43 (dt, 4.6)

3.44 (bt b, 4.8)

3.43 (dt, 4,5)

3.44 (dt, 4.6)

3.44 (t c, 4.7)

3.43 (dt, 4.5)

4.37 (bt.5.5)

2′-CH 2

3.75 (bt, 4.5)

3.73 (bt, 4.9)

3.74 (bt, 5.0)

3.73 (dt, 4.7)

3.74 (dt, 4.6)

3.74 (t, 4.6)

3.73 (dt, 4.5)

3.26 (bt, 5.5)

2-CH 2

2.38 (t, 7.4)

2.23 (t, 7.4)

2.23 (t, 7.6)

2.22 (t, 7.8)

2.21 (t, 7.7)

2.21 (t, 7.7)

2.23 (t, 7.1)

2.21 (bt, 7.3)

3-CH 2

1.59-1.66 (me)

1.66 (ttd, 7.0)

1.64 (m)

1.64 (m)

1.64 (m)

1.64 (m)

1.66 (m)

1.65 (m)

CH 2 -FAC

1.25-1.37 (m)

1.26-1.39 (m)

1.26-1.39 (m)

1.26-1.39 (m)

1.25-1.36 (m)

1.27-1.36 (m)

1.25-1.35 (m)

1.24-1.34 (m)

CH=CH-CH 2

2.04-2.08 (m)

2.01-2.05 (m)

2.00-2.03 (m)

1.93-2.00 (m)

1.98-2.05 (m)

1.94-2.00 (m)

1.99-2.07 (m)

1.95-2.02 (m)

CH=CH

5.35-5.41 (m)

5.33-5.40 (m)

5.32-5.39 (m)

5.35-5.42 (m)

5.32-5.39 (m)

5.36-5.42 (m)

5.32-5.40 (m)

5.34-5.43 (m)

CH 3

0.90 (t, 6.8)

0.90 (t, 6.4)

0.89 (t, 6.7)

0.88 (t, 6.8)

0.89 (t, 6.9)

0.88 (t, 6.9)

0.88 (t, 6.8)

0.89 (t, 6.9)

OH

n.d.f (broad)

2.78 (bs)

2.73 (bs)

2.69 (bs)

2.78 (bs)

2.3 (bs)

2.77 (bs)

NH

6.00 (bs g)

5.97 (bs)

5.98 (bs)

5.99 (bs)

5.86 (bs)

5.92 (bs)

6.00 (bs)

NH 3 +

n.d. (broad)

aDoublet of triplet

bBroad triplet

cTriplet

dTriplet of triplet

eMultiplet

fNot detected

gBroad singlet

Detection of FAEAs by UHPLC–ESI–MS/MS

To investigate if the synthesized FAEA are present in plasma from rats or humans, a multiple reaction monitoring (MRM) UHPLC–ESI–MS/MS method was developed. For this purpose, the standard substances of OEA, VEA, PeEA, EEA, tVEA, and tPeEA were fragmented separately after direct infusion. The resulting fragmentation spectra displayed high similarity (Fig. 3a) and could not be used to distinguish between the respective isomers in MRM mode, so that chromatographic separation of the substances became mandatory. Optimization of UHPLC conditions to the highest capability of the applied system resulted in clearly separated cis/trans-isomers (OEA–EEA, VEA–tVEA, and PeEA–tPeEA) as well as satisfactorily separated position isomers (VEA–OEA–PeEA and tVEA–EEA–tPeEA, Fig. 3b). EEA and PeEA co-eluted under the given chromatographic conditions preventing differentiation of these isomers. Because it was shown later that neither EEA nor PeEA were detectable in plasma from rats or humans when using the reported protocol, the co-elution of these two compounds was no major limitation for our study. The respective retention times of the separated FAEAs were assigned using the UHPLC–ESI–MS/MS parameters of the pure standards.
Fig. 3

ESI–MS/MS spectra (a) and UHPLC–ESI–MS/MS MRM (b) of the synthesized analytical standards vaccenic acid ethanolamide (VEA), oleic acid ethanolamide (OEA), trans- (t)VEA, elaidic acid ethanolamide (EEA), petroselinic acid ethanolamide (PeEA), and trans- (t)PeEA at a concentration of 1 ng mL−1. Furthermore, the signals of the 18:1 fatty acid ethanolamides (FAEAs) in rat and human plasma recorded by UHPLC–ESI–MS/MS MRM are shown

Identification and detection of VEA as a newly discovered FAEA in plasma from rats and humans

Toluene extracts of plasma samples from rats and humans were analyzed by UHPLC–ESI–MS/MS to investigate the presence of OEA, VEA, PeEA, EEA, tVEA, and tPeEA in vivo. Only VEA and OEA were detectable in rat and human plasma (Fig. 3b). The identity of both analytes was proven by the retention time and product ion scans in concentrated plasma and standard solutions. Measurement of equimolar standard samples, particularly of VEA and OEA, resulted in identical MRM response. By adding OEA-d4 as an internal standard prior to plasma extraction at a concentration of 1 ng mL−1, we were able to estimate the concentration of VEA and OEA using external calibration. In six independent rat plasma samples, the concentration of VEA ranged from 4.92 to 6.03 ng mL−1 with an average of 5.26 ± 0.57 ng mL−1, whereas OEA contents ranged from 2.38 to 3.79 ng mL−1 with an average of 3.25 ± 0.59 ng mL−1. Values for the latter analyte were in close proximity with published concentrations of 2.99 ± 0.59 ng mL−1 (n = 9) [40]. The measurement of 20 human plasma samples resulted in a range of 1.73–3.03 ng mL−1 of VEA with an average of 2.33 ± 0.43 ng mL−1 and a concentration of 3.66–10.09 ng mL−1 for OEA with an average of 5.75 ± 1.55 ng mL−1. Concentrations of OEA are higher compared to literature data with 1.37 ± 0.43 ng mL−1 (n = 23) [41] and 2.1 ± 0.1 ng mL−1 [42]. Neither PeEA nor the trans-isomers EEA, tVEA, and tPeEA could be detected in the analyzed plasma samples, although cis/trans-isomerization of fatty acids is possible in vivo [43, 44]. The absence of all FAEAs studied in this paper other than VEA and OEA could possibly be explained by the consideration that cis/trans-isomerization takes place at sites in the organism where no FAEAs are formed and hence cannot be detected in plasma. Other reasons could be that the missing FAEAs were either present in a range below the limit of detection (LOD) or that they bind to blood or plasma components and were not released during sample extraction. To explore the second possibility, we spiked human plasma samples with the respective analytes in the range of 0.05–1 ng mL−1. LOD and limit of quantification (LOQ) were then determined by calculating the signal to noise ratio (S/N) of the respective chromatograms with an S/N of 3 for LOD and an S/N of 10 for LOQ values of quantifier and qualifier transitions. The results are summarized in Table 2. Compared to LOD and LOQ values of the pure standards, represented by VEA and OEA values, LOD and LOQ in plasma are generally four times higher for the quantifier transitions and six times higher for the qualifier transitions. The recovery of internal standard OEA-d4 from human plasma matrix was 90 %. To calculate the recovery of the C18:1 isomers, the ratio of each analyte to internal standard was determined in matrix, the ratio obtained from the blank matrix was subtracted, and the resulting value compared to the ratio of the respective pure standard. The measurements were repeated three times. The recovery rate of all C18:1 isomers amounted to 90 % at a spiked concentration of 1 ng mL−1. At a spiked concentration of 0.5 ng mL−1, the recovery rate amounted to 80 % for all analytes except for tPeEA, which was at 25 %. At a spiked concentration of 0.1 ng mL−1 recovery was below 50 % for all analytes. These results are sufficient to confidently estimate the concentration of VEA and OEA in plasma, as their concentration exceeds the highest spiked plasma concentration. At the same time, our data do not allow conclusions on the contents of the other investigated endocannabinoids, which could be present below 0.5 ng mL−1 in human plasma.
Table 2

LODs and LOQs of the respective analyte transitions evaluated by plasma spiking, showing quantifier (Q) and qualifier (q) MRM transitions. See text for further information

Analyte

Retention time

Q/q

Precursor ion (m/z)

Product ion (m/z)

LOD (ng mL−1)

LOQ (ng mL−1)

VEA

7.02

Q

326.3

62.0

0.0125a

0.025a

q

326.3

95.0

0.15a

0.25a

OEA

7.12

Q

326.3

62.0

0.0125a

0.025a

q

326.3

95.0

0.15a

0.25a

tVEA

7.36

Q

326.3

62.0

0.05

0.1

q

326.3

95.0

0.5

1.5

EEA

7.46

Q

326.3

62.0

0.05

0.1

q

326.3

95.0

0.5

1

PeEA

7.43

Q

326.3

62.0

0.05

0.1

q

326.3

95.0

0.5

1

tPeEA

7.72

Q

326.3

62.0

0.05

0.1

q

326.3

95.0

1

2

aInstrumental LOD/LOQ

VEA does not form as an artifact in human and rat plasma

To our knowledge, our study was the first to separate the 18:1 isomers OEA, VEA, PeEA, EEA, tVEA, tPeEA, and, to no avail, EEA and PeEA, and to identify VEA in plasma from rats and humans.

To exclude the possibility that the VEA signal might occur as an artifact of OEA during sample preparation, e.g., by isomerization of OEA, we investigated the effects of the sample workup using an OEA standard solution. Additionally, we analyzed plasma from rats and humans with and without spiking of OEA to the plasma or blood. We could show that the workup procedure had no impact on the OEA standard solution: no peak could be observed at the retention time of VEA in this experiment. The incubation and processing of human or rat plasma after spiking with an OEA standard solution did neither lead to an increased VEA signal. The same observation was made after spiking whole blood samples of rats and humans with OEA before the plasma was gained by centrifugation. Therefore, we conclude that the signal detected in plasma samples at the retention time of the VEA standard substance did not originate from an artifact of OEA during sample preparation.

Based on the results obtained from plasma samples and standard measurements, our data strongly indicates that the novel, as yet undescribed endocannabinoid-like FAEA VEA is present in human and rat plasma besides OEA.

Possible origin of VEA in human and rat plasma

Oleic acid is highly abundant in the human diet in many common fats and oils from olive, peanut, rapeseed, or sunflower as well as in lard [15]. Additionally, meat like mutton, veal, beef, or pork contains oleic acid as the main fatty acid [15]. In contrast, vaccenic acid is only part of few and special foods like yak milk [16] or milk products like Scamorza ewe milk cheese [17]. According to the manufacturer (Altromin, Lage, Germany), the fat content of the used rat chow consists of the indigenous fat of the processed cereals and added soy oil. Thus, the main fatty acids in the chow were linoleic acid (18:2), oleic acid (18:1), and palmitic acid (16:0) with a ratio of about 4:2:1. Therefore, the high VEA content in the plasma from rats cannot be explained by a high abundance of food-derived vaccenic acid.

Besides deriving from food, vaccenic acid could originate from the metabolism of gut microbiota. It has been shown that conjugated linoleic acid and trans-vaccenic acid can be formed from linoleic acid by human fecal bacteria [45, 46] in a process which seems to be highly dependent on the resident microbiota [45]. The authors report that oleic acid (cis9-18:1) was the most abundant monounsaturated fatty acid in baseline human feces samples. After incubation with linoleic acid, gut microbiota in the samples metabolized linoleic acid, which resulted in a rise of trans-vaccenic acid levels. Other 18:1 fatty acids (like cis-vaccenic acid) are present at about one half to one third of the concentration of oleic acid [45]. Furthermore, it has been shown that vaccenic acid is mainly converted to saturated fatty acids, but also to all other 18:1 isomers in the rumen [47]. Consequently, there is evidence for an isomerization of trans-vaccenic acid to cis-vaccenic acid as well as to oleic acid and vice versa and for a variety of possible reactions triggered by different microbiota. Isomerization of oleic acid/cis-vaccenic acid was also reported to take place in higher plants [48]. It also may be speculated that OEA is converted into VEA in vivo. Probably, not only trans-vaccenic acid but also cis-vaccenic acid can be produced by gut microbiota from linoleic acid. Since we could only detect a high level of cis-VEA, but not of tVEA, the above-mentioned metabolic activities, however, remain speculative. Our analytical sensitivity might be not sufficient to detect low amounts tVEA. Besides, to the best of our knowledge, information is lacking on the enzymatic efficiency to incorporate such fatty acids into the endocannabinoid system. Consequently, all the missing fatty acids of interest may be present or formed in the gut, but could not be detected on the endocannabinoid level. More detailed investigations at the level of free fatty acids might provide better conclusions on a possible microbial origin of the described fatty acids.

A closer look at the contents of the presently detected endocannabinoids VEA and OEA in human and rat plasma, however, led to an interesting observation. Throughout all measured human samples, the ratio of OEA to VEA ranged between 2.11 and 2.77 with an average of 2.44 ± 0.27 (n = 20). In rat plasma, the ratio of VEA to OEA ranged between 1.31 and 1.93, with an average of 1.65 ± 0.22 (n = 6). Both findings suggest that the respective endocannabinoid levels in both human and rat plasma are kept constant endogenously and do not originate from ingested fatty acids. Especially the results for human plasma are striking, because we obtained random plasma samples of individuals with different eating habits and at different time points since the last meal. The constant ratios of OEA and VEA within individual plasma samples across 20 samples substantiate the assumption of an endogenous origin of VEA and OEA and their respective fatty acid precursors. Recently, studies on the elongation of long-chain fatty acids family member 6-enzyme (Elovl6) in knockout mice clarified the metabolism of fatty acid biosynthesis [49] and were later reviewed comprehensively [50]. Knockout mice deficient of Elovl6 showed an increase of C16 fatty acids, namely palmitic and palmitoleic acid, in the liver and reduced levels of C18 fatty acids like stearic, oleic, and vaccenic acid. It was also shown that Elovl6 can use palmitoleic acid as a substrate for elongation to vaccenic acid. This study also suggested that oleic acid can be synthesized by the activity of the stearoyl-CoA desaturase 1 (SCD-1) enzyme [49]. Whether vaccenic acid may be formed by SCD-1 from stearic acid and what proportions of oleic and vaccenic acid would result from this biosynthetic step cannot be deduced from these data. It appears, though, that vaccenic acid can be formed directly through a distinct biosynthetic pathway other than oleic acid. Taken together, these results suggest that mammal metabolism is able to biosynthesize oleic and vaccenic acid endogenously by means of chain elongation from shorter fatty acids and incorporate these precursors into its cannabinoid system.

Conclusion

In conclusion, we could describe the highly abundant FAEA from vaccenic acid, VEA, in plasma from rats and humans for the first time in literature. We confirmed the identity of VEA by UHPLC–ESI–MS/MS using a reference standard with the appropriate mass, fragmentation pattern, position of the double bond, and amide linkage type. We could exclude that VEA is an artifact of OEA during sample workup and discussed a possible explanation for the origin of this new metabolite of the endocannabinoid system. Due to the homology of this FAEA to the anorexigenic OEA, VEA might be associated with interesting physiological effects, which should be investigated in further studies. An intervention study with a diet abundant in palmitoleic or vaccenic acid could further clarify the metabolism of vaccinate and its derivative VEA and offer an interesting tool to explore the physiology of this newly discovered fatty acid ethanolamide.

Notes

Acknowledgments

This study is part of the Neurotrition Project, which was supported by the FAU Emerging Fields Initiative. We thank Susanne Achenbach from the blood donor bank in Erlangen, Germany, for providing human blood samples, Andreas Hess for providing the infrastructure for blood taking of rats, Johannes Niebler for his expertise regarding GC–MS measurements, and Christine Meissner for proofreading the manuscript.

Compliance with ethical standards

Conflict of interest

The authors declare that they have no conflict of interest.

Supplementary material

216_2016_9720_MOESM1_ESM.pdf (291 kb)
ESM 1 (PDF 290 kb)

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Copyright information

© Springer-Verlag Berlin Heidelberg 2016

Authors and Affiliations

  • Waldemar Röhrig
    • 1
  • Reiner Waibel
    • 2
  • Christopher Perlwitz
    • 1
  • Monika Pischetsrieder
    • 1
  • Tobias Hoch
    • 1
    Email author
  1. 1.Food Chemistry Unit, Department of Chemistry and Pharmacy, Emil Fischer CenterFriedrich-Alexander Universität Erlangen-Nürnberg (FAU)ErlangenGermany
  2. 2.Medicinal Chemistry Unit, Department of Chemistry and Pharmacy, Emil Fischer CenterFriedrich-Alexander Universität Erlangen-Nürnberg (FAU)ErlangenGermany

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