The main objectives of our study were to determine whether 5‐HT 2A receptor activation leads to the release of endocannabinoids in a mammalian expression system and, if so, to identify the signal transduction pathways responsible for this endocannabinoid production. In this report we demonstrate for the first time that the rat 5‐HT 2A receptor mediates the formation and release of the endocannabinoid 2‐arachidonoylglycerol (2‐AG) through a mechanism that is PI‐PLC dependent.

Animal models of 5‐HT 2A receptor activation employing both the 5‐HT 2A/2C ‐selective agonist DOI and cannabinoid agonists have demonstrated that 5‐HT 2A receptor‐mediated behavioral effects were negatively regulated by cannabinoid receptor 1 (CB1) activation ( Darmani 2001 ; Gorzalka et al . 2005 ). Additionally, preadministration of AM251, a CB1 antagonist, potentiated DOI‐induced wet dog shake behavior in rats. Furthermore, the endocannabinoid transport inhibitor AM404 inhibited DOI‐induced wet dog shakes in rats ( Gorzalka et al . 2005 ), suggesting a role for endocannabinoids in the DOI mechanism of action. One possible explanation for these observations involves postsynaptic 5‐HT 2A receptor activation on apical dendrites of cortical pyramidal cells, leading to endocannabinoid release, subsequent presynaptic CB1 receptor activation, and suppression of presynaptic glutamate release. This hypothetical mechanism would allow the 5‐HT 2A receptor to negatively regulate its own postsynaptic modulatory activity at excitatory synapses.

At the molecular level, the 5‐HT 2A receptor couples to G q/11 heterotrimeric G‐proteins promoting the phosphatidylinositol‐specific phospholipase C (PI‐PLC)‐mediated production of inositol triphosphate (IP3) and diacylglycerol (DAG) ( Conn and Sanders‐Bush 1986 ). The 5‐HT 2A receptor also couples directly with the monomeric G‐protein Arf1, promoting the activation of phospholipase D (PLD) and the production of phosphatidic acid (PA) ( Robertson et al . 2003 ). In addition to G q/11 and Arf1, 5‐HT 2A receptor stimulation causes activation of the monomeric G‐protein RhoA, p38 mitogen‐activated protein kinase, and p42/44 mitogen‐activated protein kinases, leading to the extracellular release of eicosanoids ( Kurrasch‐Orbaugh et al . 2003a ).

The serotonin 5‐HT 2A receptor subtype is a member of the rhodopsin‐like superfamily of G‐protein‐coupled receptors ( Hoyer et al . 2002 ). It is now widely accepted that serotonergic hallucinogens require activation of brain 5‐HT 2A receptors to produce their psychoactive effects in humans. A definitive confirmation of this mechanism is the block of the human subjective effects of the indolamine hallucinogen psilocybin by the selective 5‐HT 2A receptor antagonist, ketanserin ( Vollenweider et al . 1998 ; Carter et al . 2005 ). Not only is the 5‐HT 2A receptor involved in mediating the profound alterations in consciousness produced by hallucinogens, it is likely that the 5‐HT 2A receptor is also critical for normal cognitive functioning ( Williams et al . 2002 ). Furthermore, the most effective antipsychotic drugs also possess antagonist or inverse agonist activity at this receptor subtype ( Meltzer 1995 ).

GraphPad Prism Software (GraphPad Software Inc., San Diego, CA, USA) was used to normalize data, generate dose–response curves, and calculate intrinsic activities for 2‐AG release, AA release, PLD activation, and PLC activation. Two‐tailed paired t ‐tests were used to compare raw dpm values for inhibitor and activator treatment verses raw dpm values for 5‐HT or basal. Values were considered significantly different if they generated p < 0.05.

After treatment with m‐3M3FBS, NIH3T3–5HT 2A cells were assayed for the accumulation of inositol phosphates as previously described in detail ( Kurrasch‐Orbaugh et al . 2003b ) with the following modification: the assay was terminated after 10 min instead of the reported 30 min. As a control for PLC activation, 10 μm 5‐HT was included in the assay. Basal values averaged 123 ± 3 dpm and 5‐HT values averaged 976 ± 25 dpm for an average fold stimulation over basal of 8.

Cells were seeded into 6‐well tissue culture plates, allowed to reach 90% confluency, and incubated for 24 h prior to assay with 2.0 μCi/mL [ 3 H]palmitic acid in unsupplemented DMEM media. The cells were then washed by incubation for 15 min at 37°C with 2 mL per well of serum‐free DMEM (pH 7.4) supplemented with 0.5% fatty acid‐free BSA. Following the wash step, the media was replaced with serum‐free DMEM (pH 7.4) supplemented with 2 mL of 0.5% fatty acid‐free BSA and 0.3% n‐butanol. After 30 min treatment with agonists, cells were assayed for the accumulation of phosphatidyl butanol (PtBu). After the removal of media, the plate was placed on ice and 1 mL of ice cold MeOH/0.1 m HCl in H 2 O (1 : 1) was added to each well. Cells were scraped and transferred to glass tubes containing 1 mL ice‐cold CHCl 3 . The phases were mixed by vigorously pipetting the upper phase into the lower phase 10×. Upon separation, the lower phase was transferred to a fresh glass tube and the CHCl 3 was evaporated under a stream of argon. These sample concentrates were resuspended in 52 μL CHCl 3 and spotted onto 20 × 20 cm glass‐backed TLC plates (two 20‐μL spots/sample). The TLC plates were developed using an 11 : 5 EtOAc : isooctane mobile phase saturated with 10% (v/v) acetic acid. After drying the developed plates, the lipids were visualized with iodine vapor. The location of the phosphatidylbutanol (PtBu) spots was identified by comparison with an authentic standard and confirmed by linear radioscan. The spots were then scraped from the plates into scintillation vials, allowed to equilibrate for 3 days in scintillation fluid, and radioactivity was quantified with a liquid scintillation counter.

Several representative TLC plates were scanned using a Berthold Tracemaster 40 Automatic TLC‐Linear Analyzer (Berthold Systems Inc., Pittsburgh, PA, USA) to determine that the R f values of radioactive peaks corresponded to authentic standards of PtBu, AA, 2‐AG, and anandamide. Plates were scanned for 30–60 min and peaks were quantified using Chroma software (Berthold Systems Inc.).

The quantity of AA metabolites released was determined using a modified version of the procedure of Berg et al . (1998 ). Cells were seeded into 24‐well plates at a density of 2 × 10 5 cells/well and incubated for 24 h prior to assay with 1.0 μCi/mL [ 3 H]AA in unsupplemented DMEM media. The cells were then washed by incubation for 15 min at 37°C with 300 μL per well of serum‐free DMEM (pH 7.4) supplemented with 0.5% fatty acid‐free BSA. Each treatment condition represented four wells of a 24‐well plate. Inhibitors or antagonists were present during both this wash and the subsequent incubation with the agonist. Following removal of the wash media and addition of fresh serum‐free DMEM supplemented with 0.5% fatty acid‐free BSA and reapplication of antagonists and inhibitors when necessary, the assay was initiated by the addition of 5‐HT (10 μm), followed by incubation for 10 min at 37°C. The total volume of media per well at this stage was 500 μL, including 5‐HT and any antagonists or inhibitors. After this final incubation, 50‐μL aliquots of the cell medium were removed from each well, added to scintillation vials, and quantified using liquid scintillation counting. These data represented the total [ 3 H]AA‐labeled eicosanoids released by the cells. After removal of this aliquot, the balance of the media (1.8 mL/treatment pooled from four wells) was removed and added to a 15‐mL centrifuge tube kept on ice that contained 3.2 mL CHCl 3 , 0.8 mL MeOH, and 80 μL of 1 m HCl (40 : 10 : 1). The phases were mixed twice for 5 s with a vortex shaker and then separated by centrifugation (2000 × g , 15 min, 0°C). The CHCl 3 layer was removed from each sample and evaporated under a stream of argon. These sample concentrates were resuspended in 52 μL CHCl 3 and spotted onto 20 × 20 cm glass‐backed thin layer silica gel chromatography (TLC) plates (two 20‐μL spots/sample). The TLC plates were developed using an 13 : 5 EtOAc : isooctane mobile phase saturated with 10% (v/v) acetic acid. After drying the developed plates, the lipids were visualized with iodine vapor. Locations of the AA and 2‐AG spots were identified by comparison with authentic standards. The spots were then scraped from the plates into scintillation vials, allowed to equilibrate for 3 days in scintillation fluid, and radioactivity was quantified with a liquid scintillation counter.

[5,6,8,9,11,12,14,15– 3 H]Arachidonic acid was obtained from Amersham Life Sciences (Piscataway, NJ, USA). Bovine serum albumin (BSA) was purchased from Sigma Chemical (St. Louis, MO, USA). Ketanserin, 5‐HT, and arachidonic acid (AA) were all purchased from Research Biochemicals, Inc. (Natick, MA, USA). Anandamide, 2‐AG, AM404, MAFP, and URB597 were purchased from Cayman Chemical Co. Inc. (Ann Arbor, MI, USA). Brefeldin A, m‐3M3FBS, and D609 were purchased from Tocris Cookson Inc. (Ellisville, MO, USA). RHC80267, BAPTA‐AM, A23187, Et‐18‐OCH 3 , PMA, staurosporine, and U73122 were purchased from Calbiochem‐Novabiochem Co. (San Diego, CA, USA). Dialyzed bovine serum was purchased from Atlanta Biologicals (Lawrenceville, GA, USA). All other cell culture reagents not specifically mentioned were purchased from Gibco (Grand Island, NY, USA).

The effect of PKC inhibition and activation on 5‐HT stimulated [ 3 H]‐2‐AG release in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐AA for 24 h prior to the experiment. The effect of 5‐HT alone (over basal) was set to 100% and the treatment data are normalized to that response. The left bar illustrates the effect of 10 μm 5‐HT on [ 3 H]‐2‐AG release. The middle bar shows the effect of 100 nm staurosporine (St) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The bar on the right shows the effect of 100 nm PMA on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. Basal [ 3 H]‐2‐AG release averaged 98 ± 12 dpm and 5‐HT‐stimulated [ 3 H]‐2‐AG release averaged 218 ± 29 dpm. The data represent the mean ± SEM for at least three separate experiments. * p < 0.05 compared to 5‐HT control. ns, p > 0.05 compared with 5‐HT control.

Both calcium and DAG are produced downstream of 5‐HT 2A receptor stimulation and play a role in the activation of PKC. We have shown that DAG produced following PC‐PLC or PLD and PAH activation does not contribute to 2‐AG production in our cell line and inhibitors of this pathway lead to a small but significant stimulation of 2‐AG release over basal, while having no or a potentiating effect on 5‐HT‐stimulated 2‐AG release. It is possible that DAG produced following the activation of these enzymes leads to PKC stimulation that negatively regulates 2‐AG production or, alternatively, positively regulates 2‐AG metabolism. We thus examined the effect of PKC inhibition with 100 nm staurosporine on 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release. When NIH3T3–5HT 2A cells were pretreated with staurosporine, 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release was considerably potentiated ( Fig. 10 ). Staurosporine alone had no significant effects ( Table 2 ). Following these results, we determined the effect of the PKC activator PMA on 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release. We found that coadministration of 100 nm PMA with 10 μm 5‐HT had no significant effect on 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release ( Fig. 10 ) and PMA treatment alone had no significant effect on basal [ 3 H]‐2‐AG release ( Table 2 ). As with the inhibitors of the PLD‐PAH pathway, staurosporine had an inhibitory effect (50 ± 12%) on 5‐HT 2A receptor‐dependent [ 3 H]‐AA release, whereas PMA had no effect (95 ± 10%) on this response.

Effect of inhibitors of the PLD‐PAH pathway on 5‐HT‐stimulated [ 3 H]‐AA release in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐AA for 24 h prior to the experiment. The effect of 5‐HT alone (over basal) was set to 100% and the treatment data are normalized to that response. The leftmost bar illustrates the effect of 10 μm 5‐HT on [ 3 H]‐AA release. The 2nd bar from the left shows the effect of 100 μm brefeldin A (BFA) on 10 μm 5‐HT‐stimulated [ 3 H]‐AA release. Compare this result with the effect of 100 μm brefeldin A on PtBu accumulation ( Fig. 8 ) and 2‐AG release ( Fig. 7 ). The 3rd bar from the left shows the effect of 100 μm propranolol (Pr) on 10 μm 5‐HT‐stimulated [ 3 H]‐AA release. The rightmost bar shows the effect of 0.5% n‐butanol (Bu) on 10 μm 5‐HT‐stimulated [ 3 H]‐AA release. Basal [ 3 H]‐AA release averaged 552 ± 41 dpm and 5‐HT‐stimulated [ 3 H]‐AA release averaged 1754 ± 246 dpm. The data represent the mean ± SEM of at least three separate experiments. * p < 0.05 compared to 5‐HT control.

Primary alcohols compete for water in the second step of the PLD catalysis mechanism, inhibiting the production of PA and downstream DAG. When we included 0.5% n‐butanol (BuOH) in our assay we unexpectedly found that it had a significant potentiating effect on 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release ( Fig. 7 ). Although it is possible that the potentiating effect of BuOH may be a result of nonspecific interactions or membrane fluidity issues, 0.5% BuOH had only a slight effect on basal [ 3 H]‐2‐AG release ( Table 2 ). Nonetheless, the results mentioned above support the hypothesis that DAG generated through the 5‐HT 2A receptor‐dependent PLD pathway does not contribute to 2‐AG production, especially considering the diversity of methods used to inhibit the pathway. In the absence of 5‐HT we observed a slight but significant increase in basal [ 3 H]‐2‐AG release for all three inhibitors of the PLD pathway ( Table 2 ), a result consistent with the above hypothesis. It is possible that this pathway exerts a negative influence on 2‐AG production or alternatively a positive influence on 2‐AG hydrolysis in our cell line. Indeed, the latter case may be true as all three treatments significantly inhibited [ 3 H]‐AA release ( Fig. 9 ).

Effect of brefeldin A on 5‐HT stimulated [ 3 H]‐PtBu accumulation in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐palmitic acid for 24 h prior to the experiment. The left bar shows the basal (B) level of [ 3 H]‐PtBu accumulation. The middle bar shows the effect of 10 μm 5‐HT on [ 3 H]‐PtBu accumulation. The right bar shows the effect of 100 μm brefeldin A (BFA) on 10 μm 5‐HT‐stimulated [ 3 H]‐PtBu accumulation. Basal [ 3 H]‐PtBu accumulation averaged 340 ± 36 dpm and 5‐HT‐stimulated [ 3 H]‐PtBu accumulation averaged 3241 ± 196 dpm. The data represent the mean ± SEM for at least three separate experiments. * p < 0.05 compared to 5‐HT control.

Effect of inhibitors of the PLD‐PAH pathway on 5‐HT stimulated [ 3 H]‐2‐AG release in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐AA for 24 h prior to the experiment. The effect of 5‐HT alone (over basal) was set to 100% and the treatment data are normalized to that response. The leftmost bar illustrates the effect of 10 μm 5‐HT on [ 3 H]‐2‐AG release. The 2nd bar from the left shows the effect of 100 μm brefeldin A (BFA) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. Compare this result with the effect of 100 μm brefeldin A on PtBu accumulation ( Fig. 8 ) and [ 3 H]‐AA release ( Fig. 9 ). The 3rd bar from the left shows the effect of 100 μm propranolol (Pr) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The rightmost bar shows the effect of 0.5% n‐butanol (Bu) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. Basal [ 3 H]‐2‐AG release averaged 83 ± 4 dpm and 5‐HT‐stimulated [ 3 H]‐2‐AG release averaged 193 ± 14 dpm. The data represent the mean ± SEM for at least three separate experiments. * p < 0.05 compared to 5‐HT control. ns, p > 0.05 compared to 5‐HT control.

The effect of inhibitors of PLD‐phosphatidic acid hydrolase (PAH) pathway‐dependent DAG formation on 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release. Having demonstrated that 5‐HT‐stimulated release of [ 3 H]‐2‐AG was completely blocked by DGL inhibitors and partially blocked by the PI‐PLC inhibitor Et‐18‐OCH 3 , we wished to explore the possibility that [ 3 H]‐2‐AG release was dependent on DAG produced following 5‐HT 2A receptor‐dependent PLD and subsequent PAH activation. Therefore, cells were first pretreated with 100 μm brefeldin A, which has been reported to inhibit 5‐HT 2A receptor‐dependent PLD activity by 50% in COS‐7 cells ( Robertson et al . 2003 ). Brefeldin A had no significant effect on 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release at this concentration ( Fig. 7 ), but reduced PLD activity by 50%( Fig. 8 ). Likewise, 100 μm propranolol, reported to inhibit 2‐AG production by 50% in N18TG2 cells ( Bisogno and Di De Melck 1999 ), had no effect on 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release ( Fig. 7 ).

Effect of m‐3M3FBS, the purported PLC activator, on inositol phosphate accumulation. Cells were incubated with [ 3 H]‐myoinositol for 18 h prior to the experiment. The leftmost bar illustrates inositol phosphate accumulation following 10 min stimulation with 10 μm 5‐HT. Note that this treatment time is the same as used in all the 2‐AG release experiments. The four leftmost bars show the effect of 1–100 μm m‐3M3FBS (m‐3M3) on inositol phosphate accumulation. Compare this result with the effect of m‐3M3FBS on 2‐AG release ( Fig. 5 ). Basal (B) IP accumulation averaged 246 ± 6 dpm and 5‐HT‐stimulated IP accumulation averaged 1951 ± 49 dpm. The data represent the mean ± SEM for at least three separate experiments.

The purported PLC activator m‐3M3FBS ( Bae et al . 2003 ) stimulated a greater release of [ 3 H]‐2‐AG than 5‐HT at a concentration of 50 μm ( Fig. 5 ). As there is some controversy concerning the mechanism of m‐3M3FBS ( Krjukova et al . 2004 ), we tested the ability of a range of concentrations (1–100 μm) of m‐3M3FBS to increase the production of inositol phosphates after 10 min of stimulation. Surprisingly, but in agreement with the results of Krjukova et al . (2004 ), m‐3M3FBS did not stimulate inositol phosphate accumulation within the 10‐min time frame when 2‐AG was released ( Fig. 6 ), suggesting that m‐3M3FBS is not a direct activator of PI‐PLC but may indirectly activate PLC at a later time point following calcium elevation.

To explore the possibility that [ 3 H]‐2‐AG release was dependent on DAG produced following 5‐HT 2A receptor‐dependent PC‐PLC activation, the PC‐PLC‐specific inhibitor D609 was employed. Surprisingly, D609 at a concentration of 10 μm potentiated the 5‐HT 2A receptor‐dependent release of [ 3 H]‐2‐AG ( Fig. 5 ). A similar potentiation was observed when 100 μm D609 was used (data not shown). When D609 was examined at a concentration of 1 μm, however, the potentiating effect was not observed ( Table 2 ). We believe that the 10 μm and 100 μm concentrations of D609 are the most relevant as D609 is reported to be a competitive inhibitor of PC‐PLC with a K i of 6.4 μm ( Amtmann 1996 ).

Effects of PLC inhibition on 5‐HT‐stimulated [ 3 H]‐2‐AG release and the purported PLC activator m‐3M3FBS on [ 3 H]‐2‐AG release in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐AA for 24 h prior to the experiment. The effect of 5‐HT alone (over basal) was set to 100% and treatment data are normalized to that response. The leftmost bar illustrates the effect of 10 μm 5‐HT on [ 3 H]‐2‐AG release. The 2nd bar from the left shows the effect of 10 μm U73122 (U7) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The 3rd bar from the left shows the effect of 50 μm Et‐18‐OCH 3 (Et) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The 4th bar from the left shows the effect of 10 μm D609 (D6) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The rightmost bar shows the effect of 50 μm m‐3M3FBS (3M3) on [ 3 H]‐2‐AG release. Basal [ 3 H]‐2‐AG release averaged 82 ± 5 dpm and 5‐HT‐stimulated [ 3 H]‐2‐AG release averaged 250 ± 25 dpm. The data represent the mean ± SEM for at least three separate experiments. * p < 0.05 compared to 5‐HT control. # p < 0.05 compared with basal control.

Hypothesizing that the DAG required as a substrate by DGL was derived from 5‐HT 2A receptor‐dependent PLC activation, we employed the PLC inhibitor U73122 in our assays. U73122 at a concentration of 10 μm completely inhibited the release of [ 3 H]‐2‐AG ( Fig. 5 ). In addition, the PI‐PLC‐selective inhibitor Et‐18‐OCH 3 was employed to confirm our results with U73122. Et‐18‐OCH 3 at a concentration of 50 μm inhibited the release of [ 3 H]‐2‐AG by almost 50% ( Fig. 5 ). These results are consistent with the hypothesis that 2‐AG is produced from DAG generated by 5‐HT 2A receptor‐mediated PLC activation. Curiously, Et‐18‐OCH 3 did not fully inhibit [ 3 H]‐2‐AG release. As a possible explanation, it has been shown that Et‐18‐OCH 3 increases cytosolic calcium concentrations in Madin Darby canine kidney cells ( Jan et al . 1999 ). Considering that the calcium ionophore A23187 was able to stimulate [ 3 H]‐2‐AG release independently of 5‐HT 2A receptor activation ( Fig. 2 ), it seems possible that Et‐18‐OCH 3 partially stimulates 2‐AG release through a calcium‐dependent mechanism, while at the same time inhibiting 2‐AG release by the PLC mechanism. When examined alone, however, Et‐18‐OCH 3 failed to stimulate [ 3 H]‐2‐AG, release, although it significantly decreased basal [ 3 H]‐2‐AG release to a modest degree ( Table 2 ).

Effect of endocannabinoid transport, FAAH, and MGL inhibition on 5‐HT stimulated [ 3 H]‐2‐AG release in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐AA for 24 h prior to the experiment. The effect of 5‐HT alone (over basal) was set to 100% and treatment data are normalized to that response. The leftmost bar illustrates the effect of 10 μm 5‐HT on [ 3 H]‐2‐AG release. The 2nd bar from the left shows the effect of 10 μm AM404 (AM) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The 3rd bar from the left shows the effect of 100 nm URB597 (U597) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The rightmost bar shows the effect of 10 μm URB602 (U602) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. Basal [ 3 H]‐2‐AG release averaged 85 ± 9 dpm and 5‐HT‐stimulated [ 3 H]‐2‐AG release averaged 218 ± 19 dpm. The data represent the mean ± SEM for at least three separate experiments. * p < 0.05 compared to 5‐HT control.

After demonstrating that DGL activation was required for 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release, we examined the effect of endocannabinoid transport, FAAH, and MGL inhibition. Pretreatment with the endocannabinoid transport inhibitor AM404 at 10 μm led to a robust approximately threefold increase in 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release over 5‐HT alone ( Fig. 4 ). AM404 by itself had a slight but significant effect on basal [ 3 H]‐2‐AG release ( Table 2 ). At a higher concentration of 100 μm, AM404 increased the 5‐HT response sixfold ( Table 2 ). Similar results were obtained when the FAAH inhibitor URB597 at 100 nm was employed. The MGL inhibitor URB602 produced a modest (ca. 50%), though significant, increase in 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release at 10 μm ( Fig. 4 ). Both URB597 and AM404 had significant effects on basal [ 3 H]‐2‐AG release ( Table 2 ), although URB597 was the only inhibitor to significantly inhibit 5‐HT 2A receptor‐dependent [ 3 H]‐AA release (53 ± 8%). URB602 at 100 μm had no effect (116 ± 14%) and AM404 at 100 μm significantly potentiated (569 ± 47%) 5‐HT 2A receptor‐dependent [ 3 H]‐AA release.

Effect of DGL inhibition on 5‐HT‐stimulated [ 3 H]‐2‐AG release in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐AA for 24 h prior to the experiment. The effect of 5‐HT alone (over basal) was set to 100% and the treatment data are normalized to that response. The leftmost bar illustrates the effect of 10 μm 5‐HT on [ 3 H]‐2‐AG release. The 2nd bar from the left shows the effect of 100 μm RHC80267 (RHC) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. The 3rd bar from the left shows the effect of 1 μm THL on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release, and the rightmost bar shows the effect of 5 μm MAFP (MFP) on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. Basal [ 3 H]‐2‐AG release averaged 93 ± 10 dpm and 5‐HT‐stimulated [ 3 H]‐2‐AG release averaged 207 ± 15 dpm. The data represent the mean ± SEM for at least three separate experiments. * p < 0.05 compared to 5‐HT control.

The last step in classical 2‐AG biosynthesis is hydrolysis of the sn1 fatty acid of DAG. In the brain, this reaction is catalyzed by sn1‐specific DGLs that recently have been cloned and characterized ( Bisogno et al . 2003 ). We reasoned that inhibition of DGL should inhibit the production and release of 2‐AG. As expected, in cells prelabeled with [ 3 H]‐AA, the DGL inhibitor RHC 80267, at a concentration of 100 μm, completely inhibited the 5‐HT 2A receptor‐dependent release of [ 3 H]‐2‐AG ( Fig. 3 ). RHC 80267 failed to inhibit this response significantly at a concentration of 10 μm ( Table 2 ). Additionally, THL at 1 μm and MAFP at 5 μm inhibited 5‐HT 2A receptor‐dependent [ 3 H]‐2‐AG release by 96% and 75%, respectively ( Fig. 3 ). THL alone caused a significant reduction in basal [ 3 H]‐2‐AG release, whereas the two other DGL inhibitors failed to have any significant effect ( Table 2 ).

Effect of A23187 (A) on basal and 5‐HT‐stimulated [ 3 H]‐2‐AG release in NIH3T3‐5‐HT 2A cells. Cells were incubated with [ 3 H]‐AA for 24 h prior to the experiment. The effect of 10 μm 5‐HT alone (over basal; leftmost bar) was set to 100% and the treatment data are normalized to that response. The middle bar shows the effect of 3 μm of the calcium ionophore A23187 on [ 3 H]‐2‐AG release. The right bar shows the effect when both 3 μm A23187 and 10 μm 5‐HT are added together. Basal [ 3 H]‐2‐AG release averaged 85 ± 5 dpm and 5‐HT‐stimulated [ 3 H]‐2‐AG release averaged 236 ± 36 dpm. The data represent the mean ± SEM for at least three separate experiments. * p < 0.05 compared to 5‐HT control. # p < 0.05 compared with basal control; no significant difference from 5‐HT alone.

Release of 2‐AG stimulated by the 5‐HT 2A receptor is dose‐dependent. NIH3T3‐5‐HT 2A cells were incubated with [ 3 H]‐AA for 24 h prior to experiments. The curve illustrates one typical experiment showing [ 3 H]‐2‐AG release following 10 min stimulation with serial dilutions of 5‐HT. The inset graph illustrates the effect of 1 μm M100907 on 10 μm 5‐HT‐stimulated [ 3 H]‐2‐AG release. Basal [ 3 H]‐2‐AG release averaged 80 ± 3 dpm and 5‐HT‐stimulated [ 3 H]‐2‐AG release averaged 220 ± 29 dpm. The effect of 5‐HT alone (over basal) was set to 100% and the treatment data are normalized to that response. * p < 0.05 compared to 5‐HT control.

The 5‐HT‐stimulated release of [ 3 H]‐2‐AG was dose dependent ( Fig. 1 ) and was completely inhibited by the selective 5‐HT 2A receptor antagonist M100907 ( Sorensen et al . 1993 ; Kehne et al . 1996 ) ( Fig. 1 , inset). The 5‐HT 2A/2C selective agonist DOB, as well as the serotonergic hallucinogens 5‐MeO‐DMT, LSD, mescaline, and psilocin stimulated the release of [ 3 H]‐2‐AG in a dose‐dependent manner with EC 50 s comparable to PI hydrolysis except in the case of psilocin ( Table 1 ). Relative to total eicosanoid release, EC 50 s for 2‐AG release were higher for all agonists except LSD. The calcium ionophore, A23187 at concentrations of 3 and 6 μm, stimulated the release of [ 3 H]‐2‐AG to an extent comparable to 5‐HT ( Table 2 ). When A23187 was applied simultaneously with 10 μm 5‐HT, [ 3 H]‐2‐AG release was greater than with 5‐HT alone ( Fig. 2 ). Table 2 includes values for the response of all inhibitors and activators used in this study relative to 10 μm 5‐HT.

Three major peaks were observed by radioscanning the TLC plates. The major peak represented, on average, approximately 37% of the gross cpm for the 5‐HT lane and had an R f (0.73) corresponding to authentic AA standard. The second most abundant peak represented, on average, approximately 17% of the gross cpm for the 5‐HT lane and had an R f (0.43) corresponding to authentic 2‐AG standard. The third peak was at the origin and represented, on average, approximately 12% of the gross cpm for the 5‐HT lane. The balance of the radioactivity (about 34%) was distributed evenly along the lane and represented the background radiation. No other major peaks were observed, and a peak corresponding to anandamide was not detected. Two additional TLC solvent systems were employed to confirm that the authentic standards comigrated with peaks identified by radioscan. In diethyl ether : hexane (4 : 3) the R f s of [ 3 H]‐AA and [ 3 H]‐2‐AG were 0.34 and 0.05, respectively, and in ethyl acetate : hexane (5 : 2) the R f s were 0.50 and 0.28, respectively, corresponding to authentic standards visualized with iodine vapor.

NIH3T3–5HT 2A cells expressing the rat 5‐HT 2A receptor (5500 fmol/mg protein) averaged a threefold (two‐ to eightfold range) increase over basal in both [ 3 H]‐2‐AG and [ 3 H]‐AA release following 10 min stimulation with 10 μm 5‐HT. Recombinant A549 and HEK cell lines expressing the human 5‐HT 2A receptor (150 fmol/mg and 8000 fmol/mg protein, respectively) also exhibited similar increases over basal in both [ 3 H]‐2‐AG release and [ 3 H]‐AA release following 10 min stimulation with 10 μm 5‐HT (data not shown). The quantity of [ 3 H]‐AA released in any given assay was approximately 10 times greater than that of [ 3 H]‐2‐AG when TLC spots from the same treatment lane were quantified. The 5‐HT‐stimulated release of [ 3 H]‐2‐AG was absent in wild‐type NIH3T3 cells (data not shown), which have been reported to contain an endogenous 5‐HT 2A receptor ( Saucier and Albert 1997 ).

Discussion

In this report we have provided, for the first time, direct evidence that 5‐HT 2A receptor stimulation leads to the release of the endocannabinoid 2‐AG. Based on these experimental data, we propose that 5‐HT 2A receptor‐dependent 2‐AG release is at least partially dependent on PI‐PLC activation. We also note that DAG produced from either PC‐PLC or downstream of PLD production of PA and PAH‐mediated hydrolysis of PA does not contribute to 2‐AG biosynthesis.

For the 5‐HT 2A receptor agonists shown in Table 1, the lack of large differences between EC 50 s for IP accumulation and 2‐AG release, except in the case of psilocin, supports the hypothesis that 2‐AG release is dependent on PLC activation. Total measured eicosanoid release has been shown to be PLC independent in NIH3T3–5HT 2A cells (Kurrasch‐Orbaugh et al. 2003b). Although 2‐AG is an eicosanoid, its production was found to be PLC dependent in this same cell line. As total eicosanoid release is a measure of a population of different lipid species that are the products of multiple enzymes competing for the same AA substrate, the differences observed between the EC 50 s for 2‐AG release and total eicosanoid release are not surprising. Future studies should focus on identifying specific lipid species of the eicosanoid population that are released following 5‐HT 2A receptor activation rather than total eicosanoid release.

After determining that 2‐AG release was dependent on 5‐HT 2A receptor activation, we demonstrated that 5‐HT 2A receptor‐dependent 2‐AG release was completely inhibited by the DGL inhibitor RHC80267, which was shown by Bisogno et al. (2003) to inhibit 2‐AG production in enzyme assays using cloned DGLs. Additionally, MAFP, which is known to inhibit DGL (Moriyama et al. 1999), phospholipase A2, and FAAH, inhibited 5‐HT 2A receptor‐dependent 2‐AG release by 75%. Furthermore, 1 μm THL, which was shown by Bisogno et al. (2003) to inhibit ionomycin‐stimulated 2‐AG production in many cell types, inhibited 5‐HT 2A receptor‐dependent 2‐AG release by 96%. The endocannabinoid transport inhibitor AM404 and the FAAH inhibitor URB597 both robustly potentiated 5‐HT 2A receptor‐dependent 2‐AG release, whereas the MGL inhibitor URB602 modestly increased 5‐HT 2A receptor‐dependent 2‐AG release. Based on these data we conclude that the [3H]‐2‐AG produced following DGL activation in our cell line is metabolized by FAAH and MGL to AA and glycerol. MGL, the main 2‐AG hydrolyzing enzyme in the brain (Dinh et al. 2004; Gulyas et al. 2004; Saario et al. 2004), appears to provide less hydrolytic influence, although it remains questionable whether or not we were able to inhibit MGL fully with 100 μm U602.

We next determined that 5‐HT 2A receptor‐dependent 2‐AG release was partially dependent on DAG produced by PI‐PLC activation. The non‐selective PLC inhibitor U73122 completely inhibited 5‐HT 2A receptor‐dependent 2‐AG release at a concentration sufficient to fully inhibit inositol phosphate accumulation in NIH3T3–5HT 2A cells (Kurrasch‐Orbaugh 2002). By contrast, the PI‐PLC‐selective inhibitor Et‐18‐OCH 3 , at a concentration sufficient to inhibit inositol phosphate accumulation completely in NIH3T3–5HT 2A cells (Kurrasch‐Orbaugh et al. 2003b), reduced 5‐HT 2A receptor‐dependent 2‐AG release by only 50%. U73122 alone had no effect on basal 2‐AG release, whereas ET‐18‐OCH 3 significantly reduced basal release to a modest degree (Table 2).

When we first initiated this study, m‐3M3FBS was described as a novel nonspecific but direct PLC activator (Bae et al. 2003). Recently, however, this classification has been called into question (Krjukova et al. 2004). We therefore tested the ability of m‐3M3FBS to produce inositol phosphates in NIH3T3–5HT 2A cells. Within the 10‐min time frame when m‐3M3FBS strongly stimulated 2‐AG release, we observed no increase in the accumulation of inositol phosphates, confirming the results of Krjukova et al. (2004). Although the direct PLC stimulatory activity of m‐3M3FBS is now questionable, the stimulation of 2‐AG production may result from its effects on calcium homeostasis (Krjukova et al. 2004). Nonetheless, based on the above experiments we conclude that 5‐HT 2A receptor stimulation leads to the biosynthesis and release of 2‐AG, at least partially through a G q/11 ‐PI‐PLC‐DGL pathway.

In some cell lines, it has been shown that 2‐AG can be produced in a PLD‐dependent manner (Bisogno et al. 1999; Carrier et al. 2004). The 5‐HT 2A receptor activates PLD via the monomeric G‐protein, Arf1 (Robertson et al. 2003). PLD catalyzes the production of phosphatidylbutanol (PtBu) when stimulated in the presence of exogenous n‐butanol. Brefeldin A at 100 μm reduces PtBu production by 50% following 5‐HT 2A receptor stimulation in NIH3T3–5HT 2A cells (Fig. 8). To determine whether 2‐AG biosynthesis in our cells was dependent on DAG produced from PLD and subsequent PAH activation, we examined 2‐AG release following 5‐HT 2A receptor stimulation in the presence of brefeldin A. When NIH3T3–5HT 2A cells were treated with brefeldin A, no significant reduction in 5‐HT 2A receptor‐dependent 2‐AG release was observed. Additionally, the beta adrenergic receptor antagonist propranolol, which also inhibits PAH (Perry et al. 1992), had no effect on 5‐HT 2A receptor‐dependent 2‐AG release. When we included n‐butanol, an inhibitor of PLD‐dependent PA production, 5‐HT 2A receptor‐dependent 2‐AG release was potentiated. Based on the fact that all three of these treatments failed to inhibit 5‐HT 2A receptor‐dependent 2‐AG release, we concluded that DAG produced from PLD and subsequent PAH activation was not leading to 2‐AG biosynthesis in NIH3T3–5HT 2A cells. Curiously, all three treatments led to a significant and selective inhibition of AA release (Fig. 9) possibly by inhibiting a mechanism that positively regulates 2‐AG hydrolysis (discussed below).

Thus, we determined that at least 50% of 5‐HT 2A receptor‐dependent 2‐AG release was dependent on PI‐PLC activation but not on PLD and PAH activation. In attempts to discover the source of the remaining 50% of 2‐AG we attempted to block the response with D609, a PC‐PLC‐selective inhibitor with a reported K i of 6.4 μm (Amtmann 1996). Pretreatment with 1 μm D609 had no effect on 5‐HT 2A receptor‐dependent 2‐AG release, whereas 10 μm D609 actually led to a potentiation of 5‐HT 2A receptor‐dependent 2‐AG release. One explanation for this observation is that DAG produced following PC‐PLC activation, as with PLD/PAH activation, may lead to PKC activation, which may act either as a negative regulator of 2‐AG release or as a positive regulator of 2‐AG uptake and hydrolysis. This explanation assumes that the DAG produced following PC‐PLC activation is spatially separated from the DAG that serves as the substrate for 2‐AG biosynthesis. Indeed, the PKC inhibitor staurosporine had a similar potentiating effect on 5‐HT 2A receptor‐dependent 2‐AG release, while at the same time inhibiting AA release by 50%, as did the three PLD‐PAH pathway inhibitors. This speculation is confounded, however, by the fact that PMA had no effect on 5‐HT 2A receptor‐dependent 2‐AG or AA release. It is possible that direct activation of DGL by the 5‐HT 2A receptor through an unknown mechanism yields the other 50% of released 2‐AG, as a small pool of DAG exists in the membrane of unstimulated NIH3T3 cells (Florin‐Christensen et al. 1992). In that case, DGL activation would shift the equilibrium of upstream enzymes by reducing the pool of DAG.

A model illustrating the pathways potentially involved in the production of 2‐AG release following 5‐HT 2A receptor activation is provided in Fig. 11. In particular, the enzymatic sequences following G q/11 activation are consistent with our present results. Our experiments did not support a role for 2‐AG produced subsequent to Arf1 activation.

Figure 11 Open in figure viewer PowerPoint Model of 5‐HT 2A receptor‐dependent 2‐AG formation. The bold boxed abbreviations represent different lipid molecule pools that are modified following 5‐HT 2A receptor activation. Enzymes are presented above and below the arrows. Chemicals that modify enzymatic behavior are followed by (–) or (+) indicating the action of the chemical (inhibitory or stimulatory, respectively). Based on our results, we conclude that DAG produced following PI‐PLC activation, but not PLD‐PAH or PC‐PLC activation, is the lipid precursor for 2‐AG biosynthesis in NIH3T3‐5‐HT 2A cells.

At excitatory synapses, endocannabinoids inhibit glutamate release following postsynaptic depolarization/Ca2+‐dependent biosynthesis, retrograde transport, and activation of presynaptic CB1 receptors (Freund et al. 2003). Although the depolarization‐dependent mechanism for endocannabinoid release was documented first, there is now evidence for an additional G‐protein‐coupled receptor‐G q/11 ‐PLC mechanism (Maejima et al. 2001; Kim et al. 2002; Ohno‐Shosaku et al. 2003; Jung et al. 2005). A more recent study (Maejima et al. 2005) suggests that simultaneous depolarization and G q/11 ‐coupled G‐protein‐coupled receptor activation is more likely to be physiologically relevant than either stimulus alone. In this study, 2‐AG release was achieved by simultaneous activation of mGluR1 and depolarization with 20 mm potassium ion. Either treatment alone failed to elicit 2‐AG release. PLCβ has been shown to be the focal point where G q/11 activation and depolarization‐induced calcium influx converged to enhance endocannabinoid release (Hashimotodani et al. 2005). In our heterologous expression system, we observed an enhancement in 5‐HT 2A receptor‐dependent 2‐AG release when a calcium ionophore was included. A23187 stimulated 2‐AG release to a degree equivalent to 5‐HT 2A receptor‐dependent stimulation of 2‐AG release, and when applied simultaneously with 5‐HT we observed a response that was greater than 5‐HT alone. These results support the possibility that one of the roles of neuromodulatory neurotransmitters is to modulate depolarization/Ca2+‐dependent endocannabinoid release.

There is now abundant evidence to support the idea that endocannabinoids and neuromodulatory neurotransmitters mutually regulate the release of each other. For example, activation of dopamine (Giuffrida et al. 1999), acetylcholine (Kim et al. 2002), and metabotropic glutamate receptors (Maejima et al. 2001) has been shown to enhance endocannabinoid release. Likewise, there is evidence that endocannabinoids or CB1 agonists regulate the release of acetylcholine (Gifford and Ashby. (1996); Steffens et al. 2003), dopamine (Schlicker et al. 1996; Cadogan et al. 1997; Steffens et al. 2004), norepinephrine (Schlicker et al. 1997), and 5‐HT (Nakazi et al. 2000). It is plausible that neuromodulatory neurotransmitters, through the action of postsynaptic metabotropic receptors, partially regulate synaptic activity through the differential activation or inhibition of basal and/or depolarization‐induced endocannabinoid release.

Animal models also support a role for endocannabinoids in behaviors that result from 5‐HT 2A receptor activation. In rats, DOI reliably produces wet dog shakes, a behavioral effect mediated by 5‐HT 2A receptor activation (Gorzalka et al. 2005). Likewise in mice, DOI reliably produces both the head‐twitch response and the ear scratch response (Darmani 2001). Gorzalka et al. (2005) found that rats pretreated with the endocannabinoid uptake inhibitor AM404 displayed reduced wet dog shakes following administration of DOI. Darmani (2001) pretreated mice with cannabinoid agonists and observed both reduced head twitch response and ear scratch response when DOI was subsequently administered. By contrast, The CB1 inverse agonist SR141716A alone increased the frequency of both head twitch response and ear scratch response (Darmani and Pandya 2000).

If DOI induces the production of endocannabinoids in rats and mice through activation of the 5‐HT 2A receptor, inhibition of endocannabinoid uptake by AM404 as well as activation of CB1 receptors with CB1 receptor agonists would theoretically lead to enhanced activation of presynaptic CB1 receptors in the prefrontal cortex over that of DOI alone. Activation of presynaptic CB1 receptors would result in inhibition of presynaptic glutamate release from glutamatergic axon terminals and the reduced DOI‐induced wet dog shakes in rats and head twitch response/ear scratch response in mice that is observed. If this hypothesis is correct, preadministration of a CB1 receptor antagonist would be expected to have the opposite effect. Indeed, Gorzalka et al. (2005) found that preadministration of the CB1 receptor antagonist AM251 enhanced DOI‐induced wet dog shakes in rats.