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Endocrinology Vol. 141, No. 5 1675-1685
Copyright © 2000 by The Endocrine Society

ARTICLES

Cannabinoid CB1 Receptor-Mediated Inhibition of Prolactin Release and Signaling Mechanisms in GH4C1 Cells1

Begonia Y. Ho, Anna Stadnicka, Paul L. Prather, Arthur R. Buckley, Lori L. Current, Zeljko J. Bosnjak and Wai-Meng Kwok

Department of Pharmacology and Toxicology, University of North Dakota (B.Y.H., L.L.C.), Grand Forks, North Dakota 58203; Department of Anesthesiology, Medical College of Wisconsin (A.S., Z.J.B., W.M.K.), Milwaukee, Wisconsin 53226; Department of Pharmacology and Toxicology, College of Medicine, University of Arkansas for Medical Sciences (P.L.P.), Little Rock, Arkansas 72205; and College of Pharmacy and Department of Molecular and Cellular Physiology, College of Medicine, University of Cincinnati Medical Center (A.R.B.), Cincinnati, Ohio 45069

Address all correspondence and requests for reprints to: Dr. Begonia Ho, Department of Pharmacology, University of North Dakota, 501 North Columbia Road, Grand Forks, North Dakota 58203-2817. E-mail: bho{at}badlands.nodak.edu


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The GH4C1 cell line was used to study the cellular mechanisms of cannabinoid-mediated inhibition of PRL release. Cannabinoid CB1 receptor activation inhibited vasoactive intestinal polypeptide- and TRH-stimulated PRL release, but not its basal secretion. The cannabinoid-mediated inhibition of TRH-stimulated PRL release was reversed by the CB1 receptor-specific antagonist, SR141,716A, and was abolished by pertussis toxin pretreatment, indicating that G{alpha} subunits belonging to the Gi{alpha} and Go{alpha} family were involved in the signaling. Photoaffinity labeling using [{alpha}-32P]azidoaniline GTP showed that cannabinoid receptor stimulation in cell membranes produced activation of four G{alpha} subunits (Gi{alpha}2, Gi{alpha}3, Go{alpha}1, and Go{alpha}2), which was also reversed by SR141,716A. The CB1 receptor agonists, WIN55,212–2 and CP55,940, inhibited cAMP formation and calcium currents in GH4C1 cells. The subtypes of calcium currents inhibited by WIN55,212–2 were characterized using holding potential sensitivity and calcium channel blockers. WIN55,212–2 inhibited the {omega}-conotoxin GVIA (Conus geographus)- and {omega}-agatoxin IVA (Aigelenopsis aperta)-sensitive calcium currents, but not the nisoldipine-sensitive calcium currents, suggesting the inhibition of N- and P-type, but not L-type, calcium currents. Taken together, the present findings indicate that CB1 receptors can couple through pertussis toxin-sensitive G{alpha} subunits to inhibit adenylyl cyclase and calcium currents and suppress PRL release from GH4C1 cells.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE CANNABINOID RECEPTOR binds (-){Delta}9-tetrahydrocannabinol (THC), the active component in marijuana. In addition to its psychoactive effects, cannabinoids exhibit other physiological actions, including regulation of hypothalamic and pituitary hormones, analgesia, antiemesis, vasorelaxation, and decreasing intraocular pressure (reviewed in Refs. 1, 2). Molecular cloning has revealed two cannabinoid receptors, CB1 and CB2, that share 44% homology in their amino acid sequences (3, 4, 5). The CB1 receptor was cloned from the brain and is expressed mainly in the nervous system. The CB2 receptor was cloned from human HL60 cells and is expressed primarily in the immune system (6). The deduced amino acid sequences of both receptors indicate that they belong to the family of G protein-coupled receptors with seven putative transmembrane regions.

PRL is an anterior pituitary hormone that serves many physiological functions, including the maintenance of lactation and immunomodulation. The secretion of PRL is highly regulated by diverse neurotransmitters that act on the anterior pituitary. Its release can be stimulated by TRH through the activation of phospholipase C and subsequent activation of Ca2+ channels, as well as by vasoactive intestinal peptide (VIP), forskolin, or 8-bromo-cAMP through the stimulation of protein kinase A (7, 8, 9). Other neurotransmitters, such as dopamine, somatostatin, and acetylcholine, suppress the release by inhibiting adenylyl cyclase or attenuating calcium currents (10, 11, 12, 13). Activation of the CB1 receptor by either synthetic or endogenous ligands, such as anandamide or {Delta}9-THC, has been shown primarily to inhibit PRL secretion from the pituitary in rats (2, 14, 15) and monkeys (16), whereas the action in human remains controversial (reviewed in Ref. 2). The site where cannabinoids mediate the inhibition is also unclear. Some previous studies have indicated a direct action on the pituitary, whereas others suggest an indirect effect through the regulation of other hypothalamic neurotransmitters (2, 17).

The lactotroph-derived GH4C1 cell line and its parent rat pituitary tumor cell line, GH3 cells, have long been used to study the regulation of PRL release by numerous neurotransmitters. Previously, we reported the expression of endogenous cannabinoid CB1, but not CB2, receptors in GH4C1 cells, as identified by PCR and sequencing (18). Expression of the endogenous CB1 receptor in these cells allows us to investigate whether activation of this receptor can directly regulate PRL secretion. In addition, the GH4C1 cell has been found to express multiple G protein {alpha}-subunits as well as different effector systems, including adenylyl cyclase, phospholipase C, and Ca2+ channels (7, 19, 20, 21). Therefore, the GH4C1 cell line provides a useful model to study potential cellular mechanisms coupled to the CB1 receptor and its regulation of PRL secretion.

The signaling mechanism activated by the CB1 receptor has been studied in cell lines that express the receptor endogenously or after heterologous expression and was found to be mainly mediated through pertussis toxin (PTX)-sensitive G{alpha} subunits to produce inhibitory actions. Inhibition of adenylyl cyclase by the CB1 receptor has been reported in several cell lines, including NG108–15 and N18 cells (1, 4, 18, 22, 23). The CB1 receptor was also found to inhibit different Ca2+ channel subtypes (24, 25, 26, 27, 28, 29). In addition, membrane hyperpolarization through the activation of G protein-coupled, inwardly rectifying potassium channels (GIRK) was observed in mouse pituitary tumor AtT20 cells (29) and Xenopus oocytes (30, 31) after heterologous expression of CB1 receptors. These are all potential cellular mechanisms that may result in the inhibition of PRL release.

In the present study, we used the GH4C1 cell to test whether activation of the CB1 receptor would result in direct inhibition of PRL secretion. In addition, we examined its coupling to G proteins and two effector systems, adenylyl cyclase and Ca2+ channels.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell culture
GH4C1 cells (gift from Dr. Paul Albert, University of Ottawa, Ontario, Canada) were grown in F-10 medium containing 10% FCS, penicillin G (100 U/ml), streptomycin (100 µg/ml), and amphotericin (0.25 µg/ml). Cells were cultured in a humidified environment containing 5% CO2 and 95% air.

Quantitation of PRL release
GH4C1 cells were plated into 12-well plates the day before experiments. Cells were washed twice with PBS and preincubated with Fischer’s medium for 10 min before the addition of drugs, which were also prepared in Fischer’s medium. After a 20-min incubation, medium was collected for quantitation of the PRL released. The amount of PRL was measured by the uptake of [3H]thymidine in the PRL-dependent Nb2–11 cell, using a well established, sensitive, and highly specific lactogen bioassay (32, 33). PRL concentrations were quantitated by determining Nb2–11 cell proliferation stimulated in lactogen-free medium. Standard curves of PRL concentrations (0.01–10 ng/ml) were constructed using purified rat PRL (gift from National Hormone and Pituitary Program, lot AFP3697A). Preliminary studies showed that the concentrations of bioactive PRL determined by the Nb2–11 cell bioassay are highly correlated with those determined by RIA, and the bioassay is approximately 20-fold more sensitive (32) (our unpublished observation). The intra- and interassay coefficients of variation of the bioassay are about 5%. The amount of PRL released was normalized to the amount of cell protein measured according to the method of Bradford et al. (34).

Photoaffinity labeling
The methods for the synthesis and purification of [{alpha}-32P]azidoaniline-GTP ([{alpha}-32P]AA-GTP) and photoaffinity labeling of G{alpha} subunits with [{alpha}-32P]AA-GTP were described previously (35, 36). Briefly, GH4C1 cell membranes (100 µg/assay) were incubated in the presence or absence of agonist (10 µM WIN55,212–2 or 1 µM CP55,940) for 6 min at 30 C in 100 µl buffer I [50 mM HEPES (pH 7.4), 0.1 mM EDTA, 10 mM MgCl2, 30 mM NaCl, and 50 µM GDP]. [{alpha}-32P]AA-GTP (1 µCi/assay) was then added, and samples were incubated for an additional 10 min at 30 C. Membranes were centrifuged at 12,000 x g for 10 min and resuspended in 100 µl buffer II [50 mM HEPES (pH 7.4), 0.1 mM EDTA, 10 mM MgCl2, 30 mM NaCl, and 2 mM DTT]. Resuspended pellets were irradiated at 4 C with 240 mJ from a UV lamp (254 nm, 150 watts) at a distance of 15 cm. Samples were separated by 10% acrylamide and 6 M urea SDS-polyacrylamide gel. Proteins were transferred onto Hybond-ECL nitrocellulose membranes (Amersham Pharmacia Biotech, Arlington Heights, IL). [{alpha}-32P]AA-GTP-labeled G{alpha} subunits were visualized by autoradiography using a Molecular Dynamics, Inc., PhosphorImager 445 SI (Sunnyvale, CA) and were quantitated by densitometry using the NIH Image software program (version 1.56). To identify G{alpha} subunits, the nitrocellulose membranes were used for immunoblot analysis immediately after autoradiography, using antibodies selective for different G{alpha} subunits. Antibody-protein complexes were visualized using chemiluminescence.

cAMP assay
Adenylyl cyclase activity was assayed by cAMP formation using RIAs. Cells (0.5 x 105/assay) were preincubated with Krebs-Ringer-HEPES buffer (110 mM NaCl, 5 mM KCl, 1 mM MgCl2, 10 mM HEPES, 55 mM sucrose, and 1 mg/ml fatty-acid free BSA) containing 100 µM isobutylmethylxanthine as a phosphodiesterase inhibitor for 20 min at 37 C. Cells were then treated for 10 min at 37 C with 250 nM VIP as the adenylyl cyclase stimulator in the absence and presence of the cannabinoid receptor agonist (WIN55,212–5 or CP55,940) and antagonist (SR141,716A). The reaction was terminated by lysing the cells with trichloroacetic acid (final concentration, 5%). The lysed cells were removed by centrifugation, and the supernatant containing cAMP was ether-extracted three times. Aliquots of 50 µl were used for cAMP RIAs with anti-cAMP antibodies (gift from National Hormone and Pituitary Program, lot CV-27) according to established methods (37).

Electrophysiology
The calcium currents (ICa) in GH4C1 cells were measured using the whole cell patch-clamp technique. Cells were plated on coverslips and transferred into the recording chamber on the stage of an inverted, phase contrast microscope (IMT2, Olympus Corp., New Hyde Park, NY). The chamber was perfused at a rate of 1–2 ml/min with modified Tyrode’s solution (132 mM NaCl, 4.8 mM KCl, 1.2 mM MgCl2, 1 mM CaCl2, 10 mM HEPES, and 5 mM dextrose, with pH adjusted to 7.35 with NaOH). The internal/pipette solution contained 110 mM CsCl, 10 mM HEPES, 1 mM MgCl2, 1 mM CaCl2, 11 mM EGTA, 5 mM K2-ATP, and 0.1 mM GTP, pH 7.3 (adjusted with CsOH). Patch pipettes were pulled from borosilicate glass tubing (Garner Glass Co., Claremont, CA) using a Sachs-flaming micropipette puller (PC-84, Sutter Instrument Co., Novato, CA). The resistance of pipettes was 6–8 M{Omega}. Once the whole cell voltage clamp was established, Tyrode’s solution was replaced with a sodium-free external solution (132 mM N-methyl-D-glucamine, 4.8 mM CsCl, 5 mM dextrose, 10 mM HEPES, 2 mM MgCl2, and 10 mM CaCl2, with pH adjusted to 7.4 with HCl). WIN55,212–2 was prepared in the external solution containing 1 mg/ml fatty acid-free BSA. Currents were recorded at room temperature using a List EPC-7 patch clamp amplifier (List, Germany) interfaced to an 80486 DX computer via a TL-1 DMA interface (Axon Instruments Inc., Foster City, CA). pCLAMP software (version 6.0.2, Axon Instruments, Inc.) was used for the generation of voltage protocols, data acquisition, and analysis. The plotting program ORIGIN (version 4.1, Microcal) and Excel 97 (Microsoft Corp.) were used for additional analysis.

Currents were evoked by 50-msec depolarizing test pulses from -90 to +60 mV in 10-mV increments, from a holding potential of -90 or -50 mV. The subtracted current was determined as the difference between currents (elicited by depolarization to the same test potentials) from holding potentials of -90 and -50 mV. During whole cell recordings, the ICa had a tendency to decrease progressively. To evaluate this run-down, in a separate group of cells (n = 6), currents elicited from holding potentials of -90 and -50 mV were monitored over time under control conditions. The rate of run-down was approximately 1.5%/min. All data were corrected for the time-dependent run-down.

Determination of inositol phosphate levels
Activation of phospholipase C was estimated by the formation of inositol phosphates. The method of labeling and determination of inositol phosphate level were reported previously (31, 38). Briefly, GH4C1 cells were plated into 12-well plates. Twenty to 24 h before the experiment, the cells were labeled for 24–28 h with 1 µCi/ml myo-[2-3H]inositol (81 Ci/mmol; Amersham Pharmacia Biotech) in inositol-free DMEM (Life Technologies, Inc., Gaithersburg, MD) and 20% dialyzed FCS (Life Technologies, Inc.). The [3H]inositol-labeled cells were preincubated with 360 µl inositol-free DMEM containing 10 mM LiCl (20 min at 37 C). The reaction was started by the addition of 40 µl agonist (WIN55,212–2) in inositol-free DMEM containing 10 mM LiCl and 10 mg/ml fatty acid-free BSA, carried out for 20 min at 37 C, and terminated by rapid aspiration of the medium and the addition of 10 mM ice-cold formic acid (300 µl). After 20 min on ice, the formic acid was neutralized with 10 mM NH4OH (300 µl). Total inositol phosphates were purified from 500 µl supernatant using anion exchange resin (AG 1-X8, Bio-Rad Laboratories, Inc., Richmond, CA). Total inositol phosphate accumulation (mono-, di-, and tris-) is expressed as a percentage of the basal level determined in vehicle-treated cells in the absence of the agonist.

Statistical analysis
Data are presented as the mean ± SEM. Statistical comparisons were performed using ANOVA and paired or unpaired Student’s t test as indicated in the figure legends. IC50 values were calculated with the software Prism (version 2.0, GraphPad Software, Inc., San Diego, CA).

Materials
Synthetic {omega}-agatoxin IVA ({omega}-AgaIVA) and CP55,940 were gifts from Pfizer, Inc. (Groton, CT). Nisoldipine was a gift from Miles Pharmaceuticals (West Haven, CT). SR141,716A was a gift from Sanofi Pharmaceuticals, Inc. (Montpelier, France). WIN55,212–2, {omega}-conotoxin GVIA ({omega}-ctxGVIA, Conus geographus), and {omega}-conotoxin MVIIC ({omega}-ctxMVIIC, Agelenopis aperta) were purchased from Research Biochemicals International (Natick, MA). TRH was purchased from Peninsular Laboratories, Inc. (Belmont, CA). PTX was purchased from List Biological Laboratory (Campbell, CA). [{alpha}-32P]GTP (3000 Ci/mmol) and antisera EC2 and GC2 were purchased from NEN Life Science Products (Boston, MA). Antiserum LEP4 was a gift from Dr. Ping-Yee Law (University of Minnesota, Minneapolis, MN). Enhanced chemiluminescence reagents were purchased from Amersham Pharmacia Biotech. All other standard chemicals were purchased from Sigma-Aldrich Corp. (St. Louis, MO), Fisher Scientific (Itasca, IL), or as indicated in the text.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Inhibition of PRL release
The ability of the agonist WIN55,212–2 to inhibit PRL release was tested in GH4C1 cells. WIN55,212–2 did not inhibit the basal secretion, but significantly inhibited TRH- or VIP-stimulated PRL release (Fig. 1AGo). The inhibition of TRH-stimulated PRL release was significantly reversed by the specific CB1 receptor antagonist, SR141,716A (Fig. 1BGo), suggesting that this effect was most likely mediated by the CB1 receptor. Pretreatment with PTX abolished the inhibition (Fig. 1CGo), suggesting that G{alpha} subunits belonging to the Gi{alpha} (Gi{alpha}1, Gi{alpha}2, Gi{alpha}3) and Go{alpha} (Go{alpha}1, Go{alpha}2) families are involved in the signaling.



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  		Figure 1. Inhibition of PRL release by WIN55,212–2 from GH4C1 cells. A, GH4C1 cells were incubated with WIN55,212–2 in the absence or presence of the stimulator (1 µM TRH or 10 nM VIP). The amount of PRL released into the medium was quantitated by Nb2–11 cell bioassays as described in Materials and Methods. Results for the TRH and VIP groups are expressed as a percentage of TRH- or VIP-stimulated PRL release. Results for the no drug group are expressed as a percentage of the basal (no drug treatment) PRL level. The basal PRL level was 3.11 ± 0.52 ng/mg protein. TRH- and VIP-stimulated PRL release values were 6.30 ± 0.97 and 6.21 ± 1.45 ng/mg protein, respectively. *, P < 0.05, concentration-dependent effect for TRH- and VIP-treated cells (by ANOVA). B, Reversal of inhibition of PRL release by SR141,716A (SR). GH4C1 cells were incubated with TRH (1 µM) and WIN55,212–2 in the absence or presence of 100 nM SR141,716A. Results are expressed as a percentage of the TRH-stimulated PRL level. SR141,716A did not significantly change the TRH-stimulated PRL level (4.80 ± 2.18 ng/mg protein). **, P < 0.01, significant difference between values in the absence (-SR) and presence (+SR) of SR141,716A (by ANOVA). C, Reversal of inhibition of TRH-stimulated PRL release by pretreatment with PTX. GH4C1 cells were pretreated with PTX (50 ng/ml) for 20–24 h. Results are expressed as a percentage of the TRH-stimulated PRL level. PTX treatment did not result in a significant change in basal (3.83 ± 0.77 ng/mg protein) or TRH-stimulated (7.10 ± 1.61 ng/mg protein) PRL levels. **, P < 0.01, significant difference between values in the absence (-PTX) and presence (+PTX) of PTX (by ANOVA). The same concentrations of WIN55,212–2, TRH, VIP, and SR141,716A alone did not produce significant changes in the growth of Nb2–11 cells. All data are represented as the mean ± SEM from at least three experiments.

 
[{alpha}-32P]AA-GTP photoaffinity labeling
The identity of G{alpha} subunits activated by the CB1 receptor in GH4C1 cells was examined by measuring the increase in incorporation of [{alpha}-32P]AA-GTP produced by a maximal concentration of WIN 55,212–2 (10 µM). [{alpha}-32P]AA-GTP was incorporated into four detectable bands, designated bands i–iv, from highest to lowest mol wt (Fig. 2AGo, lane 1). The identities of the proteins that incorporated [{alpha}-32P]AA-GTP were determined by immunoblot analysis immediately after autoradiography, using antisera specific for different G{alpha} subunits: LEP4 for Gi{alpha}1 and Gi{alpha}2 (36), EC2 for Gi{alpha}3 (39), and GC2 for Go{alpha} (40).



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  		Figure 2. Photoaffinity labeling and immunoblot analysis of G{alpha} subunits activated by the CB1 receptor. A, Top panel, GH4C1 cell membranes (100 µg) were photoaffinity labeled in the presence of WIN55,212–2 (10 µM). Proteins were transferred onto nitrocellulose membranes and subjected to autoradiography (AR; lane 1) followed by immunoblotting (immunoblots, lanes 2–4) with antibodies selective for individual G{alpha} subunits: LEP4 for Gi{alpha}1 and Gi{alpha}2, EC2 for Gi{alpha}3 and GC2 for Go{alpha}1 and Go{alpha}2. Bottom panel, Autoradiogram of GH4C1 membranes (100 µg) photoaffinity labeled with no agonist, with WIN55,212–2 or WIN55,212–2 plus SR141,716A. B, Activation of multiple G{alpha} subunits in GH4C1 cell membranes treated with WIN55,212–2 or CP55,940. GH4C1 cell membranes (100 µg) were photoaffinity labeled with WIN55,212–2 or CP55,940 in the absence or presence of the CB1 receptor antagonist SR141,716A. Photoaffinity-labeled G{alpha} subunits were quantitated by densitometry. The maximum amount of G{alpha} subunits activated was defined in arbitrary OD units (the difference between the incorporation of [{alpha}-32P]AA-GTP in the absence and presence of the indicated drugs). Data represent the mean ± SEM of four to six separate experiments. Comparisons among the different G{alpha} subunits activated by each agonist were determined by paired Student’s t test (P < 0.05, a, significantly different from Gi{alpha}3; b, significantly different from Go{alpha}1; c, significantly different from Gi{alpha}2; d, significantly different from Go{alpha}2). Comparisons for the effect of each agonist in the absence and presence of SR141,716A for a given G{alpha} subunit were performed using unpaired Student’s t test (*, P < 0.05).

 
LEP4 recognized a single band (Fig. 2AGo, lane 2) that migrated in the electrophoretic mobility assay as band iii in the autoradiogram (Fig. 2AGo, lane 1). This band was designated Gi{alpha}2, because in other cell lines and tissues tested, Gi{alpha}2 migrates between Go{alpha}1 and Go{alpha}2 (35, 36), whereas Gi{alpha}1 migrates more slowly than both Go{alpha} subunits (41). EC2 recognized two major bands (Fig. 2AGo, lane 3) that migrated as the diffuse, lightly labeled autoradiographic band i (Fig. 2AGo, lane 1). As these proteins could not routinely be resolved using our urea SDS-PAGE method, autoradiographic band i was designated as the two isoforms of Gi{alpha}3 that comigrated, as previously reported in neuroblastoma x glioma NG108–15 cells (42). GC2 recognized two bands (Fig. 2AGo, lane 4) that migrated as bands ii and iv (Fig. 2AGo, lane 1), which were designated Go{alpha}1 and Go{alpha}2, respectively, as previously reported in NG108–15 cells (42). Therefore, comparison of the autoradiogram with the immunoblot indicated that the G{alpha} subunits labeled by [{alpha}-32P]AA-GTP from high to low mol wt were Gi{alpha}3, Go{alpha}1, Gi{alpha}2, and Go{alpha}2, respectively.

Stimulation of cannabinoid receptors in GH4C1 cell membranes by a maximal concentration of either WIN55,212–2 (10 µM) or CP55,940 (1 µM) produced a significant increase (P < 0.05, by paired t test) in photoaffinity labeling, relative to basal labeling, of all four G{alpha} subunits (Gi{alpha}3, Go{alpha}1, Gi{alpha}2, and Go{alpha}2; Fig. 2Go, A and B). The observed pattern of G protein activation was similar for both agonists; each produced the least amount of activation of Gi{alpha}3 (20–28 OD units), followed by slightly more stimulation of Go{alpha}1 (42–52 OD units), and the greatest activation of Gi{alpha}2 (54–80 OD units) and Go{alpha}2 (69–84 OD units). The CB1 receptor antagonist SR141,716A (1 µM) significantly attenuated activation of G{alpha} subunits stimulated by either agonist (with the exception of Gi{alpha}2 stimulation by WIN55,212–2), indicating a selective CB1 receptor-mediated effect. As Gi{alpha} and Go{alpha} are known to couple to the inhibition of adenylyl cyclase and ICa (43), these two effector systems were further investigated.

Coupling of the cannabinoid receptor to adenylyl cyclase
Stimulation of adenylyl cyclase by VIP and 8-bromo-cAMP increased PRL release from lactotrophs, whereas the inhibition of adenylyl cyclase by neurotransmitters such as somatostatin suppressed PRL release (7). Therefore, the action of CB1 receptor agonists on VIP-stimulated adenylyl cyclase was studied. WIN55,212–2 produced a concentration-dependent inhibition of VIP-stimulated cAMP formation with an estimated IC50 value of 0.43 µM (Fig. 3Go). Inhibition was also observed with CP55,940. The effects of both agonists were significantly reversed by 100 nM SR141,716A, indicating that the inhibition of adenylyl cyclase was mediated by the CB1 receptor.



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  		Figure 3. Inhibition of adenylyl cyclase by the CB1 receptor in GH4C1 cells. A, Inhibition of VIP-stimulated cAMP formation by WIN55,212–2. Cells were incubated with 250 nM VIP in the absence or presence of WIN55,212–2. Data represent a percentage of the VIP-stimulated cAMP level (n = 3–7). B, Antagonism of adenylyl cyclase inhibition by SR141,716A. Cells were incubated with 250 nM VIP and 10 µM WIN55,212–2 (WIN) or 100 nM CP55,940 (CP) in the absence (-SR) or presence (+SR) of SR141,716A (100 nM). Results are expressed as the mean ± SEM of the percentage of VIP-stimulated cAMP formation. SR alone did not result in any significant change in the VIP-stimulated cAMP level. **, P < 0.01, significantly different from -SR (by unpaired Student’s t test). Numbers in parentheses represent the number of independent experiments for each treatment. The basal and VIP-stimulated cAMP levels were 1.71 ± 0.32 and 4.21 ± 0.63 pmol/1 x 105 cells, respectively.

 
Coupling of the cannabinoid receptor to ICa
Membrane hyperpolarization is known to suppress the release of hormones and neurotransmitters. Therefore, the hypothesis that activation of the CB1 receptor may inhibit ICa was tested in GH4C1 cells. WIN55,212–2 inhibited ICa by 28.1 ± 4.2% and 23.1 ± 4.4% (Fig. 4Go) when recorded from holding potentials of -90 and -50 mV, respectively. The subtracted currents were inhibited by more than 90%. The inhibition was antagonized by SR141,716A to 1.6 ± 2.6% (Fig. 5Go). The same concentration of SR141,716A added alone did not significantly change the ICa (a change of 3.7 ± 1.8%).



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  		Figure 4. Inhibition of ICa by WIN55,212–2 in GH4C1 cells. A, Current traces of ICa recorded from a representative GH4C1 cell before (-WIN) and after (+WIN) bath application of 10 µM WIN55,212–2. Currents were recorded from a holding potential (HP) of -90 mV (left) or -50 mV (center) to a test potential of +20 mV at which peak ICa were elicited. Subtracted currents (right) were obtained by calculating the difference in currents recorded from the two holding potentials. B, Current-voltage (I-V) relationships from the same cell as that shown in A. I-V curves were constructed for currents recorded before ({circ}) and after (•) bath application of WIN55,212–2 (10 µM). Currents were elicited by test potentials of -90 to +60 mV from a holding potential of either -90 mV (left) or -50 mV (center). The I-V curves of the subtracted currents are also shown (right). C, Summary of the inhibition of ICa by WIN55,212–2. Data represent percent block of peak ICa by WIN55,212–2 normalized to peak ICa recorded in the absence of WIN55,212–2. All data were corrected for time-dependent run-down at a rate of approximately 1.5%/min as described in Materials and Methods. Numbers in parentheses represent the number of cells recorded.

 


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  		Figure 5. Antagonism of WIN55,212–2-induced inhibition of ICa by SR141,716A. Current amplitude monitored over time is shown from representative GH4C1 cells treated with WIN55,212–2 (10 µM) in the absence (A) or presence (B) of SR141,716A (200 nM). Applications of the respective drugs are indicated by bars. Currents were recorded at a test potential of +10 mV from a holding potential of -90 mV. All data were corrected for time-dependent run-down at a rate of approximately 1.5%/min.

 
Our preliminary data showed the existence of multiple Ca2+ channel subtypes in GH4C1 cells. Therefore, before determining the subtypes of ICa inhibited by WIN55,212–2, they were first characterized biophysically using holding potentials of -90 and -50 mV and pharmacologically using Ca2+ channel blockers: nisoldipine for the L-type, {omega}-ctxGVIA for the N-type, {omega}-AgaIVA for the P-type, {omega}-ctxMVIIC for the Q-type, and Ni2+ for the R-type ICa (44). The nisoldipine-sensitive ICa represented 54.4 ± 3.6% of the total ICa when recording from a holding potential of -90 mV (Fig. 6Go). This value increased to 64.3 ± 3.5% when recording from -50 mV. As the L-type ICa is not inactivated at a holding potential of -50 mV, this ICa was composed of L-type as well as residual currents through other Ca2+ channels that were not completely inactivated, such as N- and P-types. The subtracted currents between holding potentials of -90 and -50 mV would represent ICa through Ca2+ channels other than the L- type. The subtracted currents were not inhibited by the L-type channel blocker nisoldipine as expected, but were inhibited by 62.4 ± 7.3% and 90.8 ± 3.4% by {omega}-ctxGVIA and {omega}-AgaIVA, respectively (Fig. 6BGo), indicating that the subtracted currents were comprised of {omega}-ctxGVIA-sensitive N-type and {omega}-AgaIVA-sensitive P-type ICa. All GH4C1 cells tested expressed nisoldipine-sensitive L-type current; however, the {omega}-ctxGVIA-sensitive and {omega}-AgaIVA-sensitive currents were not homogeneously expressed. The blockade of the subtracted currents by {omega}-ctxGVIA ranged from 20.0–96.7%, and that by {omega}-AgaIVa ranged from 73.0–100%. This distribution explained the higher percent blockade of subtracted currents by {omega}-AgaIVA (90.8 ± 3.4%) compared with {omega}-ctxGVIA (62.4 ± 7.3%) and most likely explains the observation that the sum of ICa inhibition by both toxins exceeded 100% (Fig. 6BGo). Taken together, the nisoldipine-sensitive (L-type), {omega}-ctxGVIA-sensitive (N-type), and {omega}-AgaIVA-sensitive (P-type) currents accounted for the majority of the ICa in GH4C1 cells. No transient T-type ICa, {omega}-ctxMVIIC-sensitive Q-type ICa, or Ni2+-sensitive R-type ICa were detected in the present study (Fig. 6BGo).



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  		Figure 6. Expression of nisoldipine-, {omega}-ctxGIVA-, and {omega}-AgaIVA-sensitive ICa in GH4C1 cells. A, I-V relationships of peak ICa in GH4C1 cells before ({circ}) and after (•) bath application of nisoldipine (400 nM), {omega}-ctxGIVA (100 nM), or {omega}-AgaIVA (100 nM) from representative cells. Currents were recorded from a holding potential of -90 mV (left panel) or -50 mV (center panel). The right panel shows I-V curves of the subtracted currents. B, Summary of the inhibition of ICa by various Ca2+ channel blockers: nisoldipine (400 nM), {omega}-ctxGIVA (100 nM), {omega}-AgaIVA (100 nM), {omega}-ctxMVIIC (5 µM), and Ni2+ (50 µM). Data are presented as the percent inhibition normalized to the control peak ICa (no blocker treatment). All data were corrected for time-dependent run-down at a rate of approximately 1.5%/min. Numbers in parentheses represent the number of cells recorded in each group.

 
The Ca2+ channel subtypes that were inhibited by WIN55,212–2 were characterized by holding potential sensitivity and by the use of Ca2+ channel blockers (Fig. 7Go). At a holding potential of -90 mV, WIN55,212–2 blocked the total ICa to a similar extent before (28.1 ± 4.2%) and after (30.2 ± 4.3%) nisoldipine pretreatment (Fig. 7AGo), suggesting that WIN55,212–2 did not inhibit the L-type ICa. Subsequent to either {omega}-ctxGVIA or {omega}-AgaIVA treatment, WIN55,212–2 still inhibited the remaining ICa by only 20–30% at a holding potential of -90 mV (Fig. 7BGo), because the majority of ICa was contributed by the L-type ICa, which was not inhibited by WIN55,212–2. The subtracted currents were used to further characterize the ICa inhibited by WIN55,212–2. In control toxin-untreated cells, WIN55,212–2 inhibited the subtracted currents almost completely (92.2 ± 1.3%). In cells that were treated with {omega}-ctxGVIA to block the N-type ICa and to reveal the P-type ICa, the subtracted ICa was inhibited by WIN55,212–2 by 95.3 ± 2.0%. Similarly, in cells treated with {omega}-AgaIVA to block the P-type ICa and to reveal the N-type ICa, the subtracted ICa was inhibited by 100.0 ± 1.0%. These results suggest that WIN55,212–2 can inhibit both the {omega}-ctxGVIA-sensitive N-type and the {omega}-AgaIVA-sensitive P-type, but not the L-type ICa.



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  		Figure 7. Differential effects of WIN55,212–2 (WIN) on ICa after selective block by nisoldipine, {omega}-ctxGIVA, and {omega}-AgaIVA. GH4C1 cells were treated first with nisoldipine (400 nM), {omega}-ctxGIVA (100 nM), or {omega}-AgaIVA (100 nM) and then with WIN55,212–2 (10 µM). ICa were recorded before drug addition, after treatment with calcium channel blockers, and again after the addition of WIN55,212–2 in the presence of the indicated blocker. All data were corrected for time-dependent run-down at a rate of approximately 1.5%/min as described in Materials and Methods. Numbers in parentheses represent the number of cells recorded. A, Lack of inhibition of ICa by WIN55,212–2 after nisoldipine treatment. ICa was recorded from a -90 mV holding potential in the absence (no toxin) or presence of nisoldipine (nisoldipine-treated). WIN-sensitive current in the presence of nisoldipine was determined as the difference between the nisoldipine-treated and nisoldipine- plus WIN-treated cells. The percent inhibition by WIN55,212–2 was normalized to the total ICa recorded before the addition of nisoldipine and WIN55,212–2. B, Inhibition of ICa (holding potential, -90 mV; hatched column) and subtracted currents (open column) by WIN55,212–2 after treatment with nisoldipine, {omega}-ctxGIVA, or {omega}-AgaIVA. The percent inhibition by WIN55,212–2 was normalized to the remaining current after toxin treatment and before the addition of WIN55,212–2. Control cells were not treated with any Ca2+ channel blocker. ***, P < 0.001, the inhibition of the subtracted currents (open column) is significantly different from the inhibition of the corresponding ICa from a holding potential of -90 mV (hatched column; by one-way ANOVA after angular transformation). **, P < 0.01, the inhibition of remaining ICa in nisoldipine-treated cells is significantly different from that in the control untreated cells, both measured from -90 mV (by unpaired Student’s t test).

 
Coupling of the cannabinoid receptor to phospholipase C
Because TRH stimulates PRL release through the activation of phospholipase C (7), the effect of CB1 receptor stimulation on phospholipase C activity was investigated. GH4C1 cells have a functional phospholipase C pathway, as demonstrated by the ability of TRH (1 µM) to stimulate the formation of inositol phosphates by almost 12-fold (1197 ± 250.1% of the basal level). However, WIN55,212–2 (1 and 10 µM) did not significantly change the basal or TRH-stimulated phospholipase C activation (data not shown).


   Discussion
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The purpose of the present study was to use the GH4C1 cell as a model to study the regulation of PRL secretion by the cannabinoid CB1 receptor and its signaling mechanisms. Expression of the CB1 but not the CB2 receptor in the GH4C1 cell was first indicated by the detection of messenger RNA for the CB1 receptor and the ability of the CB1 receptor-specific antagonist, SR141,716A, to completely reverse WIN55,212-induced adenylyl cyclase inhibition (18). Moreover, activation of the CB1 receptor inhibited PRL secretion, which was also reversed by SR141,716A. This observation was consistent with the reported action of {Delta}9-THC and anandamide in inhibiting PRL release in rats (2, 14, 15) and monkeys (16), which was also antagonized by SR141,716A (14). Whether the in vivo action of cannabinoids reflects a direct effect on lactotrophs is unclear, as the hypothalamus and the pituitary have both been implicated (2, 17). Our present findings that activation of the CB1 receptor in lactotroph-derived GH4C1 cells inhibited PRL release support a direct pituitary action.

The inhibition of PRL secretion was found in the present study to be abolished by pretreatment with PTX, indicating mediation through Gi{alpha} and/or Go{alpha}. The CB1 receptor has been reported to couple to PTX-sensitive G{alpha} subunits to inhibit adenylyl cyclase and ICa (4, 24, 26, 29), but the identities of the subtypes have not yet been reported. Four subtypes of PTX-sensitive G{alpha} subunits, Gi{alpha}2, Gi{alpha}3, Go{alpha}1, and Go{alpha}2, were detected in GH4C1 cells either by immunoblot or RT-PCR (our unpublished data). Each of the four subtypes was activated by CB1 receptor stimulation, as indicated by the increases in [{alpha}-32P]AA-GTP labeling. Other endogenous receptors that inhibit PRL release in the GH4C1 cell, including the muscarinic and somatostatin receptors, have been reported to couple to multiple PTX-sensitive G{alpha} subunits, including Gi{alpha}1, Gi{alpha}2, and Gi{alpha}3 or Go{alpha} (19, 20). Although expression of Gi{alpha}1 was reported in GH4C1 cells by this group (19, 20), we were not able to detect this subunit. This apparent discrepancy may be due to the use of different medium and/or serum in culturing the cells, which may affect the transcription of different G proteins. In many cell types, receptors that interact with PTX-sensitive G proteins were found to couple to multiple effector systems, including the inhibition of adenylyl cyclase and ICa and the opening of K+ channels (43). Membrane hyperpolarization can result from both the inhibition of ICa and the opening of K+ channels. The inhibition of adenylyl cyclase may decrease cAMP formation and the phosphorylation level of Ca2+ channels by protein kinase A to cause run-down of ICa, which in turn produces membrane hyperpolarization. Inhibition of both adenylyl cyclase and ICa has been reported to suppress pituitary hormone release, including that of PRL (7). Therefore, coupling of the CB1 receptor to these potential effectors was studied systematically.

Activation of the CB1 receptor inhibited VIP-stimulated adenylyl cyclase activity, and this effect was reversed by SR141,716A. Somatostatin and dopamine have been reported to inhibit the stimulation of cAMP formation and PRL release induced by VIP, 8-bromo-cAMP, and forskolin in GH4C1 cells (7, 19). The results presented suggest that cannabinoids, dopamine, and somatostatin may suppress PRL release through a similar mechanism.

The inhibition of ICa may be the mechanism for CB1 receptor-mediated suppression of PRL release. This is supported by previous indications that activation of voltage-sensitive Ca2+ channels was found to stimulate PRL release. In addition, the inhibition of ICa by somatostatin and carbachol has been shown to suppress PRL release (10, 45). This suppression could be mediated by Gi{alpha} or Go{alpha}, as inhibition of the N- or P/Q-type ICa by the CB1 receptor has been shown to be reversed by pretreatment with PTX (24, 25, 26, 27, 28, 29).

The subtype of Ca2+ channels inhibited by the CB1 receptor varies with cell type; therefore, the subtypes affected in GH4C1 cells were investigated in detail. WIN55,212–2 inhibited the N- and P-type, but not the L-type ICa. The {omega}-ctxGVIA-sensitive N-type and {omega}-AgaIVA-sensitive P-type ICa contributed about 30–40% of the total ICa and most of the subtracted currents. The inhibition of N- and P-type ICa by WIN55,212–2 was suggested by the observation that it blocked about 30% of total ICa and almost 100% of the subtracted ICa. Similar observations were found subsequent to pretreatment of {omega}-ctxGVIA or {omega}-AgaIVA. Our findings are consistent with those of previous studies in which inhibition of the N-type, but not the L-type ICa by the endogenously expressed CB1 receptor was reported in NG108–15 and N18 cells (24, 25, 26). In AtT20 cells, activation of the heterologously expressed CB1 receptor only inhibited the Q-type not the L- or N-type ICa, even though the latter two subtypes were also present (29). In hippocampal neurons, cannabinoids inhibited both the N- and P/Q-type ICa (27). In cat cerebral arterial muscle, activation of the CB1 receptor inhibited the L-type Ca2+ channel (46). The ability of the CB1 receptor to regulate different subtypes of calcium channels could be an underlying mechanism for its broad spectrum of physiological actions.

We also studied two other effectors that may potentially mediate the action of the CB1 receptor, phospholipase C and GIRK. The observation that WIN55,212–2 did not change the basal or TRH-stimulated activity of phospholipase C indicates that the CB1 receptor-induced suppression of PRL release is not mediated through an alteration of phospholipase C activity. The potential signaling pathway through GIRK was also investigated because membrane hyperpolarization resulting from the opening of these K+ channels may suppress PRL release. In human TSH-secreting adenoma cells, somatostatin induced membrane hyperpolarization through the activation of an inwardly-rectifying K+ current via PTX-sensitive G proteins (10). The CB1 receptor has been reported to open GIRK when expressed in mouse pituitary tumor AtT20 cells and Xenopus oocytes (29, 30, 31). Carbachol and somatostatin have been shown to activate these K+ channels in the parent GH3 cells (47, 48). However, we did not detect any activation of inwardly rectifying K+ channels in GH4C1 cells exposed to WIN55,212–2 (data not shown). Carbachol (1–10 µM), somatostatin (100–500 µM), and GTP{gamma}S (500 µM) failed to open any inwardly rectifying K+ channels in these cells. Therefore, an endogenous CB1 receptor-activated, inwardly rectifying K+ channel pathway still remains to be identified.

Taken together, stimulation of the CB1 receptor resulted in the activation of PTX-sensitive Gi{alpha}2, Gi{alpha}3, Go{alpha}1 and Go{alpha}2, and the inhibition of adenylyl cyclase and N- and P-type ICa. Any of these mechanisms may participate in suppressing PRL release. As GH4C1 cells are derived from pituitary tumor cells, these results may be different in primary anterior pituitary cells. Therefore, the effects of cannabinoids on PRL secretion in primary pituitary cells are currently under investigation.

Like that of the CB1 receptor, activation of the muscarinic, somatostatin and dopamine receptors is known to inhibit PRL secretion from GH4C1 cells, GH3 cells, and pituitary glands through PTX-sensitive pathways (7, 11, 12, 49). These receptors have also been reported to signal through Gi{alpha} or Go{alpha} to inhibit adenylyl cyclase and ICa in GH4C1 cells and anterior pituitary cells (7, 8, 9, 10, 13, 45, 50). Therefore, signals elicited from different receptors may converge onto similar G protein-mediated pathways to regulate PRL release. In addition to the suppression of PRL secretion, other pharmacological actions of cannabinoids, including suppression of intraocular pressure, analgesia, and antiemesis, may be mediated by the attenuation of neurotransmission through the inhibition of ICa or adenylyl cyclase.


   Acknowledgments
 
The authors thank Jinchuan Zhao and Melissa Phelps for technical assistance, and Dr. Willis Samson for reading the manuscript. The authors acknowledge Drs. M. Parmentier and S. Munro for the complementary DNA of the CB1 and CB2 receptors, respectively.


   Footnotes
 
1 This work was supported by NIH Grants DA-09857 (to B.Y.H.), DA-10936 (to P.L.P.), and DK-53452 (to A.R.B.). Back

Received August 23, 1999.


   References
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 Introduction
 Materials and Methods
 Results
 Discussion
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