Address correspondence and reprint requests to Dr D. E. Nichols, Department of Medicinal Chemistry and Molecular Pharmacology, School of Pharmacy and Pharmacal Sciences, Purdue University, West Lafayette, IN 47907–1333, USA. E-mail: email@example.com
Previous studies in our laboratory have shown that in NIH3T3–5HT2A cells, 5-HT-induced AA release is PLA2-coupled and independent of 5-HT2A receptor-mediated PLC activation. Although 5-HT2A receptor-mediated PLC activation is known to be Gαq-coupled, much less is understood about 5-HT2A receptor-mediated PLA2 activation. Therefore, the studies presented here were aimed at elucidating the signal transduction pathway linking stimulation of the 5-HT2A receptor to PLA2 activation. By employing various selective inhibitors, toxins, and antagonistic peptide constructs, we propose that the 5-HT2A receptor can couple to PLA2 activation through two parallel signaling cascades. Initial experiments were designed to examine the role of pertussis toxin-sensitive G proteins, namely Gαi/o, as well as pertussis toxin-insensitive G proteins, namely Gα12/13, in 5-HT-induced AA release. Furthermore, inactivation of both Gβγ heterodimers and Rho proteins resulted in decreased agonist-induced AA release, without having any effect on PLC-IP accumulation. We also demonstrated 5-HT2A receptor-mediated phosphorylation of ERK1,2 and p38. Moreover, pretreatment with selective ERK1,2 and p38 inhibitors resulted in decreased 5-HT-induced AA release. Taken together, these results suggest that the 5-HT2A receptor expressed in NIH3T3 cells can couple to PLA2 activation though a complex signaling mechanism involving both Gαi/o-associated Gβγ-mediated ERK1,2 activation and Gα12/13-coupled, Rho-mediated p38 activation.
a mixture of inositol monophosphate, inositol bisphosphate, inositol triphosphate
c-Jun N-terminal kinase
mitogen-activated protein kinase
mitogen-activated protein kinase/extracellular signal-regulated kinase kinase
MAPK kinase kinase
multiplicity of infection
protein kinase C-related kinase
cytosolic phospholipase A2
sodium dodecyl sulfate–polyacrylamide gel electrophoresis
5-Hydroxytryptamine (5-HT, serotonin) receptors comprise the largest subfamily of G protein-coupled receptors cloned from mammalian tissues. Of the 14 distinct types of 5-HT receptors, classified into seven receptor classes, the 5-HT2A receptor is of particular interest to our laboratory because of its role in the action of hallucinogens and certain psychiatric illnesses. Although 5-HT2A receptors are commonly known to be Gαq-coupled, resulting in phospholipase C (PLC) activation (Hoyer et al. 1994), these receptors also can mediate stimulation of phospholipase A2 (PLA2), thereby generating the second messenger arachidonic acid (AA; Felder et al. 1990; Berg et al. 1994; Tournois et al. 1998). Having previously demonstrated (Kurrasch-Orbaugh et al. 2003) that the rat 5-HT2A receptor stably expressed in NIH3T3 cells independently activates both the PLC and PLA2 signaling pathways, i.e. that PLA2 activation was not subsequent to receptor-mediated PLC activation, we designed a series of experiments aimed at elucidating a potential mechanism by which the 5-HT2A receptor activates the PLA2 signaling cascade.
PLA2 enzymes hydrolyze the sn-2 position of membrane phospholipids to release free fatty acids and lysophospholipids. Although mammalian cells contain three structurally diverse groups of PLA2s (for a review see Capper and Marshall 2001), including Ca2+-independent PLA2 (iPLA2), secretory PLA2 (sPLA2), and 85-kDa cytosolic PLA2 (cPLA2), only cPLA2 is currently thought to be receptor-regulated (Kramer and Sharp 1997). In addition, cPLA2 has been well characterized as preferentially hydrolyzing sn-2 positions containing AA (Sharp et al. 1991; Clark et al. 1995), which is believed to be the rate-limiting step in the generation of proinflammatory lipid mediators, the eicosanoids. A wide range of receptor-mediated extracellular stimuli, including growth factors, cytokines, hormones, neurotransmitters, mitogens, antigens, and endotoxins (for a review see Murakami et al. 1997) as well as non-receptor-mediated stimuli, such as oxidation, UV light, hyperglycemia, and shear stress (Kramer and Sharp 1997), have been shown to activate cPLA2.
Regulation of cPLA2 has been studied in many different laboratories using a variety of cellular systems; hence, there exists a large-body of literature from which one can deduce differing mechanisms of activation. There does appear to be a consensus, however, that both phosphorylation of Ser505 and Ca2+ binding are critical to cPLA2 activation. Ca2+ mobilization from internal stores and/or Ca2+ influx from the extracellular space via voltage- or receptor-mediated channels, functions to translocate the enzyme from the cytosol to the membrane, where its phospholipid substrates are localized (Clark et al. 1991; Gijon et al. 1999). Stimulus-induced cPLA2 phosphorylation, however, is required for cPLA2 activity once the enzyme is localized at the membrane. Ser505 phosphorylation is commonly attributed to the mitogen-activated protein kinase (MAPK), extracellular signaling-regulated kinases (ERK1,2; p44/p42) (Lin et al. 1993; Nemenoff et al. 1993). Recent reports, however, have also demonstrated a role for p38 MAPK (Kramer et al. 1995; Geijsen et al. 2000).
The intermediary steps linking agonist-induced G-protein-coupled receptor (GPCR) stimulation to cPLA2 activation, presumably by the action of MAPKs, however, have not been elucidated. Receptor-mediated PLA2 activation has been shown to be both pertussis toxin sensitive (Gαi/o-mediated; Burch et al. 1986; Felder et al. 1990) and pertussis toxin insensitive (Burch and Axelrod 1987; Berg et al. 1998). In addition, Gβγ subunits have been shown to stimulate PLA2 activation in some cellular systems (Jelsema and Axelrod 1987; Selbie et al. 1997). Likewise, cPLA2 has been shown to be a downstream mediator of low molecular weight G proteins such as Ras and Rac, which function as switches to link biogenic-amine receptors and non-receptor tyrosine kinases to downstream effectors (Kim and Kim 1997; Yoo et al. 2001). Although there is some evidence to suggest the involvement of Gαq-coupled PLC activation in agonist-induced AA release (Lin et al. 1993), our recent results show these pathways to be regulated independently and we did not pursue further mechanisms for PLC-linked AA release.
The aim of the present study therefore was to characterize the signal transduction cascade responsible for mediating 5-HT2A receptor-coupled AA release. By employing various selective inhibitors, toxins, and antagonistic peptide constructs, we found that the 5-HT2A receptor couples to cPLA2 activation through a complex signaling mechanism involving both Gαi/o-associated Gβγ-mediated ERK1,2 activation and Gα12/13-coupled Rho-mediated p38 activation. Furthermore, inhibitors of the individual pathways failed to block completely 5-HT-induced AA release, suggesting the existence of an additional signaling component yet to be identified. Taken together, our findings elucidate potential mechanisms by which the 5-HT2A receptor stimulates AA liberation in NIH3T3–5HT2A cells.
Radioligands and chemicals
[5,6,8,9,11,12,14,15–3H]Arachidonic acid was obtained from Amersham Life Sciences (Piscataway, NJ, USA). Myo-[2-3H(N)]-inositol was obtained from New England Nuclear (Boston, MA, USA). Pertussis toxin and bovine serum albumin (BSA) were purchased from Sigma Chemical (St. Louis, MO, USA). 5-HT, ketanserin, arachidonic acid, and PD 098 059 were all purchased from Research Biochemicals, Inc. (Natick, MA, USA). SB 292 190 and SB 203 580 were obtained from BioMol (Plymouth Meeting, PA, USA). FuGENE 6 transfection reagent was purchased from Roche Diagnostics (Indianapolis, IN, USA). Dialyzed bovine serum was purchased from Hyclone (Logan, UT, USA), and all other cell culture reagents, including CRML-1066 media, were purchased from Gibco (Grand Island, NY, USA). The sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) gels and Bradford protein assay reagents were purchased from Bio-Rad (Hercules, CA, USA).
The phosphoplus p44/42 and p38 MAPK antibody kit was purchased from Cell Signaling Technology (Beverly, MA, USA). The sampler includes phospho-ERK1,2 and total ERK1,2 primary polyclonal antibodies or phospho-p38 and total p38, primary polyclonal antibodies. Anti-rabbit, Ig, fluorescein-linked whole antibody and antifluorescein alkaline phosphatase conjugate were purchased from Amersham Life Sciences (Piscataway, NJ, USA) and employed as secondary and tertiary antibodies, respectively.
The C-terminal Gβγ binding domain of βARK (Ghahremani et al. 1999) was packaged into herpes simplex virus (HSV). Likewise, the expression plasmid containing the rgRGS domain of p115RhoGEF was a kind gift from Dr Paul Sternweis (Dallas, TX, USA).
Cell culture and DNA transfection and infection
NIH3T3 fibroblasts stably expressing the 5-HT2A receptor were maintained in Dulbecco's modified Eagle's medium (DMEM), supplemented with 10% dialyzed fetal bovine serum, 2 mml-glutamine, 50 units/L penicillin, 50 µg/L streptomycin, and 300 mg/mL G-418 (all obtained from Gibco), and grown at 37°C in a 5% CO2 environment. Cells were passaged when they reached 95% confluency. Transient transfections were carried out using a 3 : 1 ratio of FuGENE 6 reagent–DNA (1.0–10 µg) in full serum media, on cells that had been plated 24 h earlier. Control assays were conducted with FuGENE 6 plus empty vector and FuGENE 6 reagent alone. The efficiency of transfection was explored using 3 : 2 and 6 : 1 FuGENE 6–DNA ratios with no additional inhibition of AA release. Phosphatidylinositol (PI) hydrolysis and AA release assays were performed after the cells had incubated for 48 h with the FuGENE 6–DNA precipitates. The C-terminal G binding domain of βARK (Ghahremani et al. 1999) was cloned into pHSVPrPUC. Replication-defective HSV vectors were packaged at a titer of approximately 2 × 105 infectious units/mL as described by Neve et al. (1997). Cells were infected with HSV-βARKCT by rinsing with PBS and incubating the cells with media containing HSV-βARKCT (approximately two infectious units/cell) 18 h prior to beginning assays. The concentration of 2.5 × 105 virions/µL was used to determine the multiplicity of infection (MOI) when 40 × 104 cells/well were seeded.
Cells were grown on six-well plates and subjected to the same procedure as employed for PI hydrolysis and AA release assays. Following a 30-min incubation with or without 5-HT, the cells were lysed [2 mm Hepes, 1 mm EDTA, 0.3 mm phenylmethylsulfonyl fluoride (PMSF), 1 mm dithiothreitol (DTT), 100 mm Na3VO4, pH 7.4], homogenized, and centrifuged. The supernatant was removed and the pellet resuspended (15 mm Hepes, 0.3 mm PMSF, 1 mm DTT, 1 mm Na3VO4, pH 7.4). A Bradford protein assay was conducted to quantify protein and ensure equal loading. Protein samples were heated at 100°C for 5 min and subjected to SDS–PAGE on 4–15% acrylamide gels, followed by transfer to polyvinylidine difluoride membranes for 3 h. Blocking of membranes occurred overnight at 4°C in Tris-buffered saline (TBS) with 5% (w/v) non fat dried milk. The membranes were then rinsed and incubated with the primary antibody in TBS for 3 h at room temperature, followed by secondary and tertiary antibody incubations for 1 h each, in TBS supplemented with 0.5% Tween-20. The blots were developed using enhanced chemifluorescence kits (ECF, Amersham Life Sciences) and results were obtained using the Storm™ fluorescence-scanning instrument (Molecular Dynamics, Sunnyvale, CA, USA).
PI hydrolysis assays
Accumulation of total IP was determined using a modified version of a previously published protocol (Berg et al. 1994). Cells were seeded in 48-well plates at a density of 1 × 105 cells/well and labeled 18 h prior to assay by the addition of 1.0 µCi/mL myo-[2-3H(N)]-inositol in serum-free CRML-1066 media. For the assay, the cells were pretreated for 15 min with 10 µm pargyline, 10 mm LiCl, and any inhibitors, and then were stimulated with agonists for 30 min at 37°C. The assay was terminated by aspirating the medium and adding 10 mm formic acid. [3H]Phosphoinositides were separated on a Dowex-1 ion exchange column (Berridge et al. 1983), eluted with 1.0 m ammonium formate and 0.10 m formic acid, and quantified using liquid scintillation counting. The range of basal [3H]IP accumulation ± treatment was 3–5% of maximal 5-HT stimulation. For example, maximal 5-HT-mediated (10 µm) stimulation of [3H]IP accumulation was 7000–10 000 cpm, compared with basal levels of 250–500 cpm.
The quantity of released AA 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 × 105 cells/well and labeled with 0.5 µCi/mL [3H]AA for 5 h prior to assay. After this incubation, the cells were washed by a 15-min incubation with serum-free DMEM supplemented with 0.5% fatty acid-free BSA in a 37°C water bath. Inhibitors or antagonists were present during the wash. The assay was initiated by the addition of 5-HT (10 µm) followed by incubation for 30 min at 37°C. After that final incubation, an aliquot of the cell medium was removed, added to scintillation vials, and quantified using liquid scintillation counting. The range of basal [3H]AA release ± treatment was 18–22% of maximal 5-HT stimulation. For example, maximal 5-HT-mediated (10 µm) [3H]AA release was 2000–2500 cpm, with basal levels of 400–550 cpm.
The effect of Gαi/o and Gα12/13 inhibition on PLA2-AA release and PLC-IP accumulation
Because our previous results (Kurrasch-Orbaugh et al. 2003) had demonstrated 5-HT2A-coupled AA release to be (i) PLA2-dependent (presumably cPLA2) and (ii) independent of receptor-mediated PLC activation, we next wished to explore the involvement of specific signaling molecules in agonist-induced AA release, in an attempt to elucidate the signaling cascade responsible for linking 5-HT2A receptor activation to PLA2 stimulation in NIH3T3–5HT2A cells.
Initial experiments were conducted to explore the role of Gαi/o-family G proteins in 5-HT2A receptor-mediated AA release. This family of G proteins was selected because some laboratories have shown GPCR-mediated PLA2 activation to be pertussis toxin sensitive (Burch et al. 1986; Felder et al. 1990), even though others have found it to be pertussis toxin insensitive (Burch and Axelrod 1987; Berg et al. 1998). Pertussis toxin is a bacterial toxin that functions to ADP-ribosylate Gαi, Gαo, and Gαt proteins, thereby preventing the activation of these proteins by GPCRs (Katada and Ui 1982).
Pretreatment with pertussis toxin (PTX; 25 ng/mL) for 18 h partially inhibited 5-HT-mediated AA release when compared with control (63 ± 4.6% decrease; Fig. 1). In contrast, PTX pretreatment had no effect on PLC-IP accumulation (100 ± 5%; Fig. 1). In an attempt to obtain a further increase in PTX-inhibited AA release, the concentration of PTX was incrementally increased from 25 ng/mL to as high as 150 ng/mL, which is equal to or higher than the concentrations of PTX used by other laboratories studying Gαi/o-mediated signaling in NIH3T3 cells (Gupta et al. 1990; Sa and Das 1999). Even at the highest concentration of PTX(150 ng/mL) 5-HT-mediated AA release was inhibited only 64 ± 5.7%, and the PTX-insensitive Gαq-coupled PLC-IP accumulation remained unaffected (100 ± 11%).
The inability of PTX to block 5-HT2A receptor-mediated PLA2 activation beyond ca. 50% suggests that in addition to activation of a Gαi/o-coupled signaling cascade, the 5-HT2A receptor also may be coupling to a PTX-insensitive signaling pathway to activate PLA2. Possible signaling proteins that may link 5-HT2A receptor activation to PLA2 stimulation include the PTX-insensitive G proteins Gαs, Gαq, and Gα12/13, adapter proteins including β-arrestins and G protein receptor kinases (GRKs), and trans-activation of receptor tyrosine kinases. Several lines of evidence led to the selection of Gα12/13 proteins as potential candidates that might link 5-HT2A receptors to AA release. For example, Gα12/13 has been shown to potentiate serum-induced PLA2 activation (Xu et al. 1993) as well as to regulate COX-2 (the enzyme responsible for the conversion of free AA into prostaglandins) activity (Dermott et al. 1999). In addition, Gα12/13-coupled MAPK activation has been reported (Dhanasekaran and Dermott 1996; Sah et al. 2000), and MAPK have been shown to phosphorylate, and subsequently activate PLA2 (Lin et al. 1993).
To explore the consequences of Gα12/13 inhibition for 5-HT-induced PLA2-AA release and PLC-IP accumulation, an antagonistic peptide, RhoGEF Regulator of G protein Signaling (rgRGS), was employed. The rgRGS domain, located outside the RGS box of p115RhoGEF, selectively binds to Gα12/13 G proteins but confers no GAP activity (Chen et al. 2001). Thus, we employed the rgRGS construct to function as a Gα12/13-selective inhibitor to explore the role of Gα12/13 in 5-HT2A receptor-mediated PLA2-AA release. Following transient transfection with rgRGS, 5-HT-induced PLA2-AA release was partially inhibited (47 ± 2.3% decrease; Fig. 2), whereas there was no effect on PLC-IP accumulation (110 ± 4.4%; Fig. 2). In an attempt to increase the rgRGS-mediated inhibition of 5-HT-induced AA release, the concentration of DNA was increased 5-fold (to 5.0 µg), which resulted in a decrease of PLA2-AA release (29 ± 6.0%), although there was still no effect on PLC-IP accumulation at this higher concentration (108%, n = 1).
Because the above results demonstrate that 5-HT2A receptor-mediated AA release is partially inhibited by PTX-sensitive Gαi/o (63 ± 4.6% decrease; Fig. 1) and also by rgRGS-sensitive Gα12/13 (47 ± 2.3% decrease; Fig. 2), additional studies were conducted to determine whether treatment with both PTX and rgRGS would result in 100% inhibition of 5-HT-coupled AA release. Surprisingly, no additional inhibition of 5-HT-induced AA release was observed in NIH3T3–5HT2A cells exposed to PTX + rgRGS (52 ± 1.5% decrease), compared with cells treated with each inhibitor alone. Furthermore, there was no effect on PLC-IP accumulation in cells treated with both PTX and rgRGS (105 ± 5.5%). Taken together, these data suggest that in NIH3T3–5HT2A cells, 5-HT2A receptor-mediated PLA2-AA release is both Gαi/o- and Gα12/13-coupled. The lack of an additive effect following combined treatment with PTX + rgRGS was, however, unexpected. Nevertheless, the ability of rgRGS to inhibit 5-HT-mediated AA release, at least partially, suggests a role for Gα12/13.
The effect of Gβγ sequestration and Rho inhibition on PLA2-AA release and PLC-IP accumulation
Having demonstrated a potential role for Gαi/o and Gα12/13 in receptor-mediated PLA2 activation, additional studies were conducted to identify the downstream signaling pathway responsible for linking 5-HT2A receptor-Gαi/o and -Gα12/13 activation to AA release. Initial studies were aimed at exploring the role of the Gβγ heterodimer for two reasons. First, Gαi/o–Gβγ–Ras–Raf–MEK–ERK activation has been well documented (Hawes et al. 1995), and ERK has been shown to phosphorylate PLA2 (Lin et al. 1993; Nemenoff et al. 1993). Second, a role for Gβγ in agonist-induced PLA2 activation has already been demonstrated in bovine rod outer segments (Jelsema and Axelrod 1987).
Thus, to obtain further insight into the role of Gβγ subunits in 5-HT-mediated AA release in NIH3T3–5HT2A cells, a Gβγ-scavenger was employed to function as an antagonistic peptide by binding free Gβγ subunits and preventing their interaction with effectors. In particular, the Gβγ binding domain located at the C-terminal end of the β-adrenergic receptor kinase (βARKCT; Ghahremani et al. 1999), was cloned into the herpes simplex virus (HSV) to facilitate viral-mediated gene delivery of this construct into NIH3T3–5HT2A cells. Following infection with HSV-βARKCT (2.0 MOI) for 18 h, 5-HT-mediated PLA2-AA release was partially inhibited (50 ± 8% decrease), whereas this treatment had no effect on PLC-IP accumulation (100 ± 9.3%; Fig. 3). To examine expression levels following infection, western blot analysis for HA-βARKCT expression was employed following 0, 0.5, 1.0, and 2.0 MOI; the expression of βARKCT was dose-dependent and was not detected in mock infected cells (data not shown).
Further, to test the hypothesis that the Gβγ subunits were originating from a PTX-sensitive heterotrimeric G protein, NIH3T3–5HT2A cells were simultaneously exposed to HSV-βARKCT and PTX (25–100 ng/mL) for 18 h. If the Gβγ dimer was originating from a heterotrimeric G protein other than Gαi/o, then pretreatment with PTX + βARKCT should result in complete inhibition of 5-HT-mediated AA release because the individual treatments with these inhibitors each resulted in c. 50% inhibition of AA release. In contrast, if Gαi/o and Gβγ were originating from the same heterotrimeric G protein, then pretreatment with PTX + βARKCT should not be additive because inhibition of either Gα or Gβγ would block components of the same signaling pathway, and only ca. 50% inhibition would be expected. Following exposure of NIH3T3–5HT2A cells to PTX + βARKCT, only 45 ± 6.3% inhibition of PLA2-AA release was observed, which is nearly equal to the extent of AA release observed when the cells are treated with PTX or βARKCT alone. Furthermore, this treatment had no effect on PLC-IP accumulation. Thus, these data suggest that 5-HT2A receptor-mediated PLA2 activation is in part coupled to Gαi/o and Gβγ, which possibly are originating from the same heterotrimeric G protein. Consistent with our hypothesis, these results suggest that pretreatment with PTX partially inhibits PLA2 activation by preventing the release of Gβγ from activated Gαi/o-family G proteins.
Because the above results suggest that Gαi/o and Gβγ are functioning in a linear pathway, additional studies were conducted to explore the role of Gα12/13 and its downstream effectors, namely Rho proteins, in 5-HT-induced AA release. Even though the Rho gene family is composed of 14 members, broadly divided into four subfamilies (Rho, Rac, Cdc42, and Rnd; Sah et al. 2000), only the Rho subfamily, consisting of RhoA, RhoB, and RhoC, was considered. Furthermore, although the C3 toxin is a selective Rho subfamily inhibitor that does not distinguish between RhoA, RhoB, and RhoC, the more general terminology of Rho will be employed.
Gα12/13-coupled Rho activation has been demonstrated in several cell lines (Hart et al. 1998; Katoh et al. 1998) including NIH3T3 cells (Tolkacheva et al. 1997; Mao et al. 1998). Thus, additional studies were conducted to explore the effects of Rho inactivation on 5-HT-mediated AA release in NIH3T3–5HT2A cells. In particular, C3 transferase, a toxin that specifically ribosylates Asn41 within the Rho effector-binding site, was employed to inactivate Rho by preventing interaction with downstream substrates (Sekine et al. 1989). Expression of the C3 toxin in NIH3T3–5HT2A cells resulted in a partial inhibition (48 ± 4.3% decrease; Fig. 4) of 5-HT-mediated PLA2-AA release, whereas this toxin had no effect on PLC-IP accumulation (106 ± 3.0%; Fig. 4). In an attempt to determine whether 5-HT-coupled Gα12/13 activation and 5-HT-coupled Rho activation were occurring in a linear pathway, NIH3T3–5HT2A cells were simultaneously pretreated with both the Gα12/13-specific rgRGS construct and the Rho-specific C3 transferase. If Gα12/13 and Rho were functioning in a linear signaling pathway, then their combined effect on 5-HT-induced AA release should be no greater than the effect produced by each inhibitor alone. Interestingly, following a 48-h incubation with NIH3T3–5HT2A cells, 5-HT-mediated AA release was inhibited only 25 ± 6.0%, less than the inhibition observed when these vectors were expressed separately. Taken together, these data suggest that Rho proteins may in part be responsible for 5-HT-directed AA release, and that 5-HT2A receptor-directed Rho stimulation may be Gα12/13-coupled.
Demonstration of 5-HT-mediated ERK1,2 and p38 activation
cPLA2 is regulated by Ca2+ binding and by phosphorylation. Although Ca2+ is necessary for cPLA2 activity because it mediates translocation of the enzyme to the membrane where its phospholipid substrates reside, Ca2+ binding is not sufficient for cPLA2 activation (Clark et al. 1991). Instead, activation of cPLA2 requires phosphorylation. In particular, the extracellular signaling-regulated kinases (ERK1,2; p44/p42; Lin et al. 1993; Nemenoff et al. 1993) and the p38 MAP kinases (Kramer et al. 1995; Geijsen et al. 2000) both have been shown to phosphorylate cPLA2. Therefore, additional studies were conducted to explore the role of ERK1,2 and p38 in 5-HT2A receptor-mediated PLA2-AA release in NIH3T3–5HT2A cells. Because 5-HT2A receptor activation leads to stimulation of MAPKs in large arteries (Watts 1996), smooth muscle (Kelleher et al. 1995), and aortic tissues (Florian and Watts 1998), we hypothesized that incubation with 5-HT would result in activation of ERK1,2 and p38 MAPKs in our cellular system.
Phospho-ERK1,2 and phospho-p38 selective antibodies were employed to determine the effect of receptor activation on ERK1,2 and p38 stimulation. As predicted, when NIH3T3–5HT2A cells were exposed to 5-HT (10 µm) for 30 min, a significant increase in ERK1,2 (Fig. 5) and p38 (Fig. 6) phosphorylation was observed. This increase was abolished by pretreatment with the 5-HT2A receptor antagonist, ketanserin (10 µm; Figs 5 and 6), demonstrating that ERK1,2 and p38 stimulation was receptor-mediated.
Pretreatment with the MEK1,2 inhibitor PD 098 059 (10 µm), partially blocked 5-HT-mediated ERK1,2 phosphorylation (Fig. 5). MEK has not been found to have kinase activity for any substrate other than ERK, thus inhibition of MEK by PD 098 059 results in selective inhibition of the ERK1,2 signaling cascade (Mansour et al. 1996). Taken together, these data show that agonist-induced stimulation of the 5-HT2A receptor in NIH3T3–5HT2A cells leads to ERK1,2 and p38 activation.
The effect of ERK1,2 and p38 inhibition on PLA2-AA release and PLC-IP accumulation
Having shown that 5-HT can stimulate ERK1,2 and p38 MAPK phosphorylation in NIH3T3–5HT2A cells, it was important to demonstrate that inhibition of these kinases resulted in decreased 5-HT2A receptor-mediated PLA2-AA release. To determine the potential role of ERK1,2 MAPK in 5-HT-induced PLA2 activation, the MEK-selective inhibitor PD 098 059 was employed. Following pretreatment of NIH3T3–5HT2A cells with PD 098 059 (1.0 µm and 10 µm) a partial inhibition of 5-HT-mediated PLA2-AA release was observed (42 ± 3.9% and 54 ± 3.6% decreases, respectively; Fig. 7), whereas there was no effect on PLC-IP accumulation (93 ± 2.0% and 97 ± 6.5%, respectively; Fig. 7). Timecourse experiments demonstrated that 15 min was the optimal preincubation time for both ERK and p38 inhibitors. Increasing the concentration of PD 098,059 to 30 µm gave no additional PLA2-AA inhibition (35 ± 6.1% decrease; solubility problems arose at concentrations higher than 30 µm).
To explore the potential role of p38 in 5-HT2A receptor-induced PLA2 activation, SB 202,190, the p38-selective inhibitor, was employed. Crystal structure and mutagenesis analysis have been used to demonstrate the specificity of pyridinyl imidazole inhibitors such as SB 202 190 for p38 MAPK over JNK or ERK (Wilson et al. 1997). Thus, incubation with SB 202 190 should selectively target p38 MAPK and decrease 5-HT-induced AA release if p38 is functioning as a 5-HT2A receptor-coupled downstream signaling molecule. Pretreatment of NIH3T3–5HT2A cells with SB 202 190 (1.0 µm and 10 µm), partially inhibited 5-HT-mediated PLA2-AA release (19 ± 4.2% and 55 ± 4.3% decrease, respectively; Fig. 8), although PLC-IP accumulation remained unaffected (103 ± 2.3% and 102 ± 2.1%, respectively; Fig. 8). Increasing the concentration of SB 202,190 to 30 µm provided no additional inhibition of AA release (31 ± 7.3% decrease), nor was there any effect on PLC-IP accumulation (100 ± 0.50%; solubility problems arose at concentrations higher than 30 µm).
In an attempt to produce a larger inhibition of 5-HT-mediated PLA2-AA release than that observed with SB 202,190, we employed SB 203,580, another pyridinyl imidazole p38 inhibitor similar to SB 202,190, at a peak concentration of 50 µm. No further inhibition was observed (36 ± 3.3% decrease). Finally, the combined inhibition of ERK by PD 098 059 and p38 by SB 202 190 was not additive, i.e. 5-HT-induced AA release was not completely blocked. Taken together, these data suggest that receptor-induced PLA2-AA release is mediated, at least in part, by the ERK1,2 and p38 MAPK signaling cascades.
In the present study, we showed that the 5-HT2A receptor couples to PLA2-AA release through a complex signaling cascade that involves multiple G proteins and the ERK1,2 and p38 MAPKs in NIH3T3 cells. Our previous studies (Kurrasch-Orbaugh et al. 2003) revealed that 5-HT2A receptor-mediated AA release was PLA2-dependent but PLC-independent, because PLC, DAG lipase, and protein kinase C (PKC) inhibitors had no effect on 5-HT2A receptor-mediated AA release. Thus, the studies conducted herein were designed to map out a potential signaling cascade linking 5-HT2A receptor activation to PLA2-AA release.
Several approaches were taken to show that multiple G proteins and intracellular signaling molecules might in part mediate 5-HT-induced PLA2-AA release. The data suggest that the 5-HT2A receptor may couple to PLA2 activation through at least two parallel signaling cascades. For example, pretreatment with PTX partially abolished AA release, suggesting a role for Gαi/o in 5-HT-induced AA release. Furthermore, infection with the Gβγ-scavenger, HSV-βARKCT, resulted in ca. 50% inhibition of 5-HT-mediated AA release, thereby also implicating Gβγ. Finally, pretreatment with the MEK inhibitor, PD 098,059, resulted in a significant decrease of 5-HT2A receptor-mediated AA release when compared with control cells. Therefore, taken together, these data map out a potential signaling cascade linking stimulation of the 5-HT2A receptor to PLA2-AA release: activation of the pertussis toxin-sensitive G protein, namely Gαi/o, causes the release of Gβγ, which is free to initiate activation of the Ras–Raf–MEK–ERK signaling cascade, ultimately leading to ERK-mediated phosphorylation of cPLA2 (see Fig. 9).
Because each of the inhibitors employed along this Gαi/o-coupled signaling pathway resulted only in ca. 50% inhibition of 5-HT-induced AA release, additional studies were conducted to explore the existence of a second 5-HT2A receptor-mediated signaling pathway. Transient expression of a Gα12/13-selective inhibitor, rgRGS, attenuated 5-HT-induced AA release, suggesting a role for Gα12/13 in agonist-mediated AA release. In addition, in the presence of C3 transferase, which specifically targets Rho, 5-HT2A receptor-mediated PLA2 activation was significantly decreased, thereby also implicating a role for Rho in receptor-driven AA release. Finally, incubation with the p38-selective inhibitor, SB 202,190, resulted in partial inhibition of 5-HT-induced AA release. Taken together, these data suggest that 5-HT2A receptor-coupled PLA2 activation may be linked through an additional signaling cascade. We propose that the second signaling pathway may be mediated by receptor-coupled pertussis toxin-insensitive Gα12/13, which functions to activate Rho, and ultimately results in p38-mediated phosphorylation of cPLA2 (see Fig. 9).
Even though corroborative reports from the literature substantiate a role for each of these signaling molecules in 5-HT2A receptor-mediated AA release, the organization of these signaling molecules into specific, linear signaling cascades appears to be inaccurate; ultimately, the nature of these cascades may not be so systematic. As such, the signaling molecules shown here to mediate receptor-induced AA release may or may not be functioning within the limits of the proposed pathways. For example, it is proposed that 5-HT2A receptor activation results in PLA2 stimulation through a Gαi/o–βγ–Ras–Raf–MEK–ERK signaling cascade. Nevertheless, agonist-induced p38 activation also has been shown to be pertussis toxin-sensitive as well as Gβγ-mediated (Communal et al. 2000; Yamauchi et al. 2001), thereby making Gαi/o–Gβγ–p38 an equally likely signaling pathway to link 5-HT2A receptor activation to PLA2 stimulation. Furthermore, Rho has been shown to enhance synergistically Raf-mediated activation of ERK1,2, thus providing modulation of ERK1,2 activation independent of the proposed Gαi/o–βγ–Ras–Raf signaling cascade (Luttrell et al. 1997). Likewise, activated Ras can stimulate Rho proteins in a Raf-independent manner, which suggests a potential Gαi/o–βγ–Ras–Rho–p38 signaling pathway that links 5-HT2A receptor activation to a Rho–p38 coupled singling cascade (Zohn et al. 1998). Finally, although receptor-mediated signaling is often G protein–dependent, recent results suggest that other intracellular signaling proteins, such as β-arrestins, GRKs, and proteins that contain motifs known to modulate protein–protein interactions (i.e. SH2, SH3, PDZ domains) may associate directly with the receptor, thereby serving as a scaffold with which additional signaling proteins may interact to form a complex responsible for activation of signaling cascades, independent of heterotrimeric G proteins (Hall et al. 1999). Thus, although the signal transduction pathways proposed in Fig. 9 are well substantiated in the literature, additional studies are needed to establish accurately the manner in which Gαi/o, Gα12,13, Gβγ, Rho, ERK, and p38 signaling molecules mediate 5-HT2A receptor-coupled PLA2 activation in NIH3T3–5HT2A cells. Further, it should be noted that one caveat to studies that employ MAPK inhibitors is that they can be non-selective, making conclusions about the exact role of p38 and p42/44 in 5-HT2A receptor-mediated AA release difficult.
One of the most striking results in the data obtained from this study is the inability of any of the inhibitors, toxins, or constructs employed (alone or in combination) to abolish completely 5-HT-induced AA release. In fact, only the 5-HT2A receptor antagonist ketanserin and the PLA2 inhibitor mepacrine were able to inhibit all 5-HT-induced AA release in NIH3T3–5HT2A cells (Kurrasch-Orbaugh et al. 2003). One explanation of these findings is that both of the pathways studied share a common enzymatic ‘rate limiting’ step prior to PLA2 activation. As discussed above, ERK1,2 and p38 are possible candidates for this role. Thus, maximal activation of either of the G proteins we have implicated ultimately proceeds through this common enzymatic step, which itself is not capable of fully activating PLA2. There appears to be no precedent, however, for enzymatic mechanisms that would lead only to partial activation of PLA2. Perhaps a more likely possibility is that other members of the PLA2 enzyme family, namely sPLA2, could be mediating agonist-induced liberation of AA independent of Gαi/o- and Gα12/13-coupled PLA2 (presumably cPLA2) activation. Even though cPLA2 is often considered to be the isozyme responsible for stimulus-induced eicosanoid production, several recent reports have shown that sPLA2 activation also contributes to prostaglandin production (Fonteh et al. 1994; Murakami et al. 1996; Murakami et al. 1998), suggesting that both cPLA2 and sPLA2 may modulate AA metabolism. Furthermore, sPLA2 is not regulated by postreceptor signaling such as Ca2+ binding and phosphorylation; therefore, activation of sPLA2 would probably be through a signaling mechanism that is independent of 5-HT2A receptor-coupled cPLA2 activation. In addition, evidence exists to suggest that cPLA2 and sPLA2 can interact with each other. Conflicting reports have resulted in two alternative mechanisms, one that proposes that sPLA2 can activate cPLA2 (Hernandez et al. 1998) and one that proposes that cPLA2 contributes to sPLA2 activation (Balsinde and Dennis 1996). Nonetheless, bi-directional interaction between cPLA2 and sPLA2 may represent an additional component to 5-HT-induced AA release apart from our proposed signal transduction pathway leading to cPLA2 activation.
In conclusion, these data provide evidence that 5-HT2A receptor-coupled PLA2 activation involves a complex signaling paradigm. In particular, 5-HT-induced AA release was shown to be both Gαi/o- and Gα12/13-mediated. Furthermore, a role for Gβγ and Rho proteins in agonist-induced PLA2 activation was demonstrated. Finally, as hypothesized, ERK1,2 and p38 MAPKs were shown to be activated in NIH3T3–5HT2A cells following stimulation with 5-HT. Likewise, inhibition of these kinases resulted in a decrease of 5-HT2A receptor-activated AA release, thereby suggesting that cPLA2 activation is dependent on MAPK-coupled phosphorylation. Taken together, it is proposed that the 5-HT2A receptor couples to PLA2 activation, at least in part, by activating two parallel signaling pathways, one that is comprised of Gαi/o–Gβγ–Ras–Raf–ERK1,2 and another that is Gα12/13–Rho–p38.
In addition, because inhibition of either pathway provided a nearly identical reduction in AA release, it seems likely that the two pathways share a common final enzyme prior to PLA2 activation. These data are exciting because they serve to outline the mechanism by which 5-HT2A receptors stimulate AA mobilization in NIH3T3–5HT2A cells. Furthermore, in terms of agonist-directed trafficking, these results show that 5-HT2A receptor-mediated PLA2 activation is considerably more complex than 5-HT2A receptor-mediated PLC activation. This observation raises the possibility that cellular responses mediated by 5-HT2A receptor activation may arise from stimulation of other complex signaling cascades, beyond the robust Gαq-directed PLC activation that is the most easily measured response to 5-HT2A receptor activation.
The authors would like to thank Dr Paul Albert (McGill University, Montreal, Canada) for providing us with the βARKCT construct and Timothy Vortherms for cloning the βARKCT into herpes simplex virus (Purdue University; West Lafayette, IN, USA). In addition, we thank Clark Wells and Dr Paul Sternweis (University of Texas-South-western Medical Center, Dallas, TX, USA) for the construction and generous donation of the rgRGS expression plasmid. This work was supported by National Institutes of Health Grant DA02189.