Corticotropin-Releasing Hormone-Mediated Induction of Intracellular Signaling Pathways and Brain-Derived Neurotrophic Factor Expression Is I
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内分泌学杂志 2005年第3期
Department of Pathobiochemistry (N.B., C.B.), Johannes Gutenberg University Mainz, 55099 Mainz, Germany; and Group Molecular Genetics of Behaviour (H.H., B.L.), Max Planck Institute of Psychiatry, 80804 Munich, Germany
Address all correspondence and requests for reprints to: Dr. Christian Behl, Department of Pathobiochemistry, Johannes Gutenberg University Mainz Medical School, 55099 Mainz, Germany. E-mail: cbehl@uni-mainz.de.
Abstract
CRH receptor (CRHR) 1 and the cannabinoid receptor 1 (CB1) are both G protein-coupled receptors. Activation of CRHR1 leads to increases in cAMP production and phosphorylation of the transcription factor cAMP response element-binding protein (CREB). In contrast, CB1 is negatively coupled to the cAMP signaling cascade. In this study, we analyzed a putative interaction between these two systems focusing on the regulation of the expression of brain-derived neurotrophic factor (BDNF), a CREB-regulated gene. In situ hybridization revealed coexpression of CRHR1 and CB1 receptors in the granular layer of the cerebellum. Therefore, we analyzed the effects of CRH and the CB1 agonist WIN-55,212-2 on BDNF expression in primary cerebellar neurons from rats and mice. We observed that application of CRH for 48 h led to an increase in BDNF mRNA and protein levels. This effect was inhibited by WIN-55,212-2. At the level of intracellular signaling, short-term application of WIN-55,212-2 inhibited CRH-induced cAMP accumulation and CREB phosphorylation. Pharmacological analysis demonstrated that the CRHR1 antagonist R121919, the protein kinase A inhibitor H89, and the calcium chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid acetoxymethyl ester inhibited CRH-mediated BDNF expression. Moreover, depolarization-induced BDNF synthesis was also inhibited by long-term application of WIN-55,212-2 in wild-type mice but not in CB1-deficient mice. Thus, these data highlight an interaction between the CRH and the cannabinoid system in the regulation of BDNF expression by influencing cAMP and Ca2+ signaling pathways.
Introduction
CRH, A MAJOR mediator of the stress response in the central nervous system (CNS) (1) also affects other central processes, such as learning and memory, synaptic plasticity, and neuroprotection (2, 3, 4). These effects are mediated by CRH binding and activation of two distinct G protein-coupled receptors, CRH receptor (CRHR) 1 and CRHR2, which are found throughout the CNS and the periphery (5). CRH has a higher affinity for CRHR1 than for CRHR2, and in the brain, CRHR1 is expressed at high levels in the hippocampus, cortex, and cerebellum (6). CRH binding to CRHR1 typically activates adenylate cyclase (AC), which leads to increased intracellular concentrations of cAMP and activation of protein kinase A (PKA) (5). However, downstream target genes of CRH have not yet been thoroughly investigated; previous studies have focused on genes involved in the stress response and ACTH release, such as proopiomelanocortin (7). One putative target is the brain-derived neurotrophic factor (BDNF), the expression of which is controlled by cAMP-elevating agents in neurons (8). In addition to its role as a classical target-derived growth factor during neuronal development, BDNF is an essential autocrine factor, released and acting locally after neuronal depolarization (9).
Cannabinoids also exert most of their effects through activation of G protein-coupled receptors, which include the inhibition of AC through Gi/o proteins (10). To date, two cannabinoid receptors have been identified; the cannabinoid receptor type 1 (CB1) is widely expressed throughout the CNS (11), whereas the cannabinoid receptor type 2 is mainly present in immune cells (12). Among other regions, CB1 is localized densely in the cerebellum of the postnatal and adult brain (13, 14).
A striking feature of the cannabinoid system is its involvement in the modulation of various neurotransmitter systems, mainly by decreasing neurotransmitter release (15). Despite the recent progress in understanding the actions of endocannabinoids on synaptic transmission (16), the signal transduction pathways regulated by Gi/o-coupled CB1 in the cerebellum are poorly characterized. Most of the data were obtained in nonneuronal cell lines, in which stimulation of CB1 activates the ERK subtype of MAPK (17) eventually inducing the expression of the immediate-early genes coding for c-fos and Zif268 (18). Although there is evidence that activation of CB1 by exogenous ligands (9-tetrahydrocannabinol and WIN-55,212-2) regulates the release of ACTH via the secretion of CRH within the hypothalamic-pituitary-adrenal (HPA) axis (19, 20), signaling interactions between the corresponding receptors and subsequent effects on downstream transcriptional targets have not been investigated in detail. Intriguingly, a rapid component of glucocorticoid-negative feedback, the process that controls the stress response by affecting the HPA axis through the inhibition and chronic reduction of expression of CRH in the hypothalamus, has been recently reported to be mediated in a cannabinoid/CB1-dependent manner (21).
In the present study, we investigated other putative modes of interaction between the cannabinoid system and the CRH system regarding intracellular signaling mechanisms. As a model system, we used primary cultures of cerebellar granule neurons where both receptors are highly expressed. Cerebellar granule cells constitute the largest homogeneous neuronal population of mammalian brain. Due to their postnatal generation and the feasibility of well-characterized primary in vitro cultures, cerebellar granule neurons have acquired a special position in neuroscience as one of the most reliable models for the study of neural development, function, and pathology. We examined the effects of CRH and the CB1 agonist WIN-55,212-2 on BDNF expression, the transcription of which is reported to be dependent on cAMP response element-binding protein (CREB) phosphorylation and neuronal activity and might, therefore, represent a downstream target.
Materials and Methods
Drugs
3-Isobutyl-1-methylxanthine (IBMX; Sigma, Deisenhofen, Germany), WIN-55,212-2 (Tocris, Cologne, Germany), rat/human CRH (Calbiochem, Schwalbach, Germany), 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid acetoxymethyl ester (BAPTA-AM) (Calbiochem), H89 (Calbiochem), and R121919 (kind gift from Dr. F. Ohl, Max Planck Institute of Psychiatry, Munich, Germany) were prepared as concentrated stock solutions in PBS (IBMX and H89, both 10 mM), 100% dimethylsulfoxide (DMSO; WIN-55,212-2, 10 mM, and BAPTA-AM, 1 mM), and 2% acetic acid (CRH, 10 mM) or citrate buffer (R121919, 100 mg/ml).
Animals and tissue preparation
Sprague Dawley rats, C57BL/6N mice (both from Charles River, Sulzfeld, Germany), CB1-deficient mice, and wild-type littermates (22) were housed with a 12-h light, 12-h dark cycle and allowed to access food and water ad libitum. All experimental protocols were approved by the Ethical Committee on Animal Care and Use of the Governments of Bavaria and Rheinland-Palatinate, Germany.
Adult mice (3–5 months old) were killed by cervical dislocation for in situ hybridization experiments, and newborn rats or mice were decapitated for cerebellar primary cultures. Brains were snap frozen on dry ice immediately after isolation and stored at –80 C until sectioning. Brains were mounted on Tissue Tek (Polysciences, Warrington, PA), and 18-μM-thick coronal consecutive sections were cut from the cerebellum on a cryostat (Microtome HM 560; Microm, Braunschweig, Germany). Sections were mounted onto frozen SuperFrost/plus slides (Fisher Scientific, Schwerte, Germany), dried on a 42 C warming plate, and stored at –20 C until used.
Synthesis of probes
35S-labeled and digoxigenin (DIG)-labeled riboprobes were used. CRHR1 plasmid was a kind gift from Dr. W. Wurst (Max Planck Institute of Psychiatry) and was used as described previously (23). The CB1 probe was used as described in Marsicano and Lutz (24). In vitro transcription was carried out for 35S-labeled and DIG-labeled riboprobes, as previously described (25). Using these probes for in situ hybridization experiments, sense controls did not give any detectable signals, and antisense probes gave expression patterns identical to those already published in rat or mouse (data not shown).
In situ hybridization
Single in situ hybridization with 35S-labeled riboprobes for CB1 and CRHR1 was carried out as previously described (24), and double in situ hybridization was carried out as previously described (25). The TSA Biotin System (NEN Life Science Products, Boston, MA) was used for detection of the DIG-labeled CB1 probe, and the chromogenic reaction was carried out with the Vector Red kit (Vector Laboratories, Gruenberg, Germany). Slides were dipped in photographic emulsion (NTB-2 from Kodak, Rochester, NY; diluted 1:1 in distilled H2O) for detection of the 35S-labeled CRHR1. After 4 wk of exposure at 4 C, slides were developed (D-19; Kodak) and fixed (Kodak fixer).
Cell culture
Primary rat or mouse cerebellar granular cultures were prepared as described previously with modifications (26). Cerebellar tissue from 3-d-old animals was dissected, and tissue pieces were incubated for 20 min in Ca2+-free, Mg2+-free Dulbecco’s PBS (Invitrogen, Karlsruhe, Germany) containing 0.1% trypsin and 0.02% EDTA (pH 8.0) at room temperature. Cells were transferred to Ca2+-free, Mg2+-free Hank’s balanced salt solution (Invitrogen) supplemented with 10% fetal calf serum (Invitrogen) and dissociated gently by pipetting up and down. Undissociated pieces were filtered through a 50-μM pore-sized Nybolt mesh (Eckardt, Waldkirch, Germany), and cells were centrifuged at 200 x g for 4 min. The pellet was resuspended in MEM (Invitrogen) supplemented with 10% horse serum (Invitrogen), and the number of viable cells was counted by trypan blue exclusion. Cells were seeded (150,000 cells/cm2) into poly-L-ornithine-coated (0.1 mg/ml; molecular mass, 100–200 kDa; Sigma) 6- or 24-well plates with MEM (Invitrogen) supplemented with 10% horse serum in the case of rat cultures or MEM supplemented with 20% fetal calf serum in the case of mouse cultures. Culture medium was changed to serum-free N2-supplemented MEM/F12 (Invitrogen) medium after 24 h for rat cultures and Neurobasal medium supplemented with B27 (both from Invitrogen) for mouse cultures. Cells were used for experiments after another 24 h.
cAMP accumulation assay
The cAMP assay was performed as described previously (27) with slight modifications. Primary cultures were maintained in 24-well plates, and IBMX was added to cultures 5 min before addition of CRH (10–8 M) and/or WIN-55,212-2 (10–6 M). Cells were incubated for 10 min with the drugs, and reactions were terminated by aspiration of the medium and addition of 1 ml ice-cold 6% trichloroacetic acid, followed by incubation overnight at 4 C. Two percent acetic acid alone and 0.2% DMSO used as vehicle controls had no effect on cAMP accumulation. The extracts were treated twice with 4 ml diethylether to remove the trichloroacetic acid, dried overnight in a lyophilisator, and reconstituted in serum-free culture medium. Intracellular cAMP levels were measured with a competitive protein binding assay (NEN Life Science Products). Data are expressed as percentage of basal cAMP levels. Extracts from two wells were pooled, and samples were measured in triplicates.
Immunoblot analysis
Cells plated in six-well dishes were lysed in 200 μl lysis buffer (60 mM Tris/HCl, pH 8.0; 2% sodium dodecyl sulfate, 10% sucrose, 5 μg/ml aprotinin, and 10 mM phenylmethylsulfonyl fluoride) per well. Lysates were briefly sonicated and stored at –80 C. Protein content was measured using the BCA assay (Pierce, Bonn, Germany). Samples were diluted 1:1 in 2x sample buffer (250 mM Tris-HCl, pH 6.8; 4% sodium dodecyl sulfate, 10% glycerol, 2% ?-mercaptoethanol, and 0.0006% bromophenol blue) and denatured at 95 C for 5 min. Proteins (10 μg/lane) were separated by SDS-PAGE (10% resolving gel) and transferred onto a nitrocellulose membrane (Schleicher and Schuell, Dassel, Germany) by semidry transfer (Bio-Rad, Munich, Germany). Blots were blocked for 30 min at room temperature with blocking buffer (5% fat-free milk powder, 0.05% Tween 20, 20 mM Tris/HCl, pH 7.6; and 150 mM NaCl) and incubated with either anti-CREB (1:500; Calbiochem) or anti-pCREB (phosphorylated at serine 133; Upstate Biotechnologies, Lake Placid, NY) primary antibodies in blocking buffer overnight at 4 C. After incubation with a horseradish peroxidase-coupled secondary antibody for 1.5 h (1:5000; Dianova, Hamburg, Germany) at room temperature, bands were visualized using the ECL detection system (Pharmacia, Freiburg, Germany). ODs of bands were calculated with Scion software (Scion Corp., Frederick, MA). Phosphorylated protein levels were normalized to total unphosphorylated levels.
Semiquantitative RT-PCR
Total RNA was isolated from cultures with peqGOLD RNAPure (Peqlab, Erlangen, Germany) according to the manufacturer’s instructions. Residual genomic DNA was removed with RNase-free DNase I. RNA (5 μg) was used for Superscript II (Life Technologies, Karlsruhe, Germany) reverse transcriptase-mediated synthesis of oligo(dT)12–18-primed (Roche, Mannheim, Germany) cDNA. PCR was carried out as follows: 94 C for 1 min; 55 C for hypoxanthinguanine phosphoribosyl transferase (HPRT) and ?-actin or 63 C for BDNF for 1 min; and 72 C for 1 min, with a 10-min extension at 72 C during the last cycle. PCR was carried out with 28 and 32 cycles for HPRT/?-actin and BDNF, respectively. Primer sequences included the following: BDNF (28); HPRT: sense 5'-CCTGCTGGATTACATTAAAGCACTG-3'; antisense 5'-GTC AAG GGC ATA TCC AAC AAA C-3'; and ?-actin: sense 5'-CTA CAA TGA GCT GCG TGT GGC-3'; antisense 5'-CAG GTC CAG ACG CAG GAT GGC-3'. PCR products of 297, 351, and 275 bp for BDNF, HPRT, and ?-actin, respectively, were amplified. Negative RNA controls without reverse transcriptase ensured a lack of genomic DNA contamination. ODs of PCR bands were measured with the Kodak-1D software. Results were calculated as ratios of OD of the BDNF vs. HPRT or ?-actin bands, respectively. To ensure that the PCR products fell within the linear range, cycle dependency was carried out for BDNF, HPRT and ?-actin. PCR samples were run on agarose gels, and ODs were measured using Scion Image software (Scion Corp.) and plotted on graphs. In each case, the correlation coefficient was r2 0.97 (data not shown).
ELISA for BDNF expression
The Emax immunoassay system (Promega, Mannheim, Germany) was used to quantify the levels of BDNF protein in primary neuronal cultures. DMSO (0.2%) vehicle control had no effect on BDNF expression. Lysis of cells, determination of protein content, and ELISA procedure were carried out as described in detail elsewhere (29). Samples from three pooled wells were measured in duplicates.
Statistical analysis
Data were analyzed by one-way ANOVA using GraphPad (GraphPad Prism, San Diego, CA) or SigmaStat (Systat Software, Point Richmond, CA) software. Significance between groups was further analyzed using the post hoc Tukey test, and data were always depicted with SEM. P < 0.05 was considered statistically significant.
Results
Colocalization of CB1 and CRHR1 in the mouse cerebellum
Both CB1 and CRHR1 receptors have been described to be expressed in the postnatal and adult cerebellum. In adult mouse and rat cerebellum, CRHR1 receptor mRNA and protein was described to be present throughout the cerebellum in Purkinje cells, the granule cell layer, and within all four of the cerebellar nuclei (30). Binding studies in early postnatal mice (postnatal d 3) showed CRHR1 distribution throughout the cerebellum. At postnatal d 10, the adult pattern of distribution and level of labeling begins to emerge (31). Similarly, CB1 was detected in the postnatal rat cerebellum by binding studies, in situ hybridization and immunohistochemistry showed an increasing signal density in adulthood, where it is mainly expressed throughout the granule and molecular layers (13, 14). However, the locations of both CB1 and CRHR1 within the same cells must be demonstrated to put forward a putative interaction between these two receptor systems.
In Fig. 1, we show two sections of the cerebellum that were hybridized with 35S-labeled riboprobes for either CB1 (Fig. 1A) or CRHR1 (Fig. 1B), indicating the high expression levels of both receptors in this layer. To show that both receptors are indeed present in the same cells, we performed double in situ hybridization by combining a DIG-labeled riboprobe for CB1 with a 35S-labeled riboprobe for CRHR1. Due to the high density of cells in the granule layer, it was not feasible to count coexpressing cells at a single-cell resolution in this area. Because silver grains (expression of CRHR1) and red staining (expression of CB1) are uniformly distributed throughout all layers of granule cells, we estimated a 100% coexpression of CB1 and CRHR1 receptors (Fig. 1C).
FIG. 1. Bright- and dark-field micrographs of coronal cerebellar sections showing examples of coexpression of CB1 with CRHR1 as detected by single or double in situ hybridization, respectively. Expression of (A) CB1 and (B) CRHR1 in the cerebellum as detected with 35S-labeled riboprobes. Scale bars, 1 mm. C, Coexpression of CB1 (red staining) and CRHR1 (silver grains) in the granule layer. Scale bar, 50 μm. ML, Molecular layer; GL, granular layer.
CRH induces BDNF expression in cerebellar cultures through calcium- and PKA-dependent signaling pathways
The majority of the actions of CRH have been demonstrated to be mediated through the activation of AC and the subsequent elevation in intracellular cAMP levels (5). Moreover, CRH-mediated ACTH release in corticotrope cells has been shown to involve L-type Ca2+ channels (32). Elevations in intracellular cAMP or activation of Ca2+-dependent pathways result in the induction of BDNF expression (8, 33). Therefore, we examined the effects of CRHR1 activation on BDNF expression in cerebellar cultures. Semiquantitative RT-PCR analysis was carried out using BDNF-specific primers on cDNA derived from cells stimulated with CRH alone or in combination with the PKA inhibitor H89 (10 μM), the calcium chelator BAPTA-AM (1 μM), and the CRHR1 antagonist R121919 (100 ng/ml), respectively. Relative expression was determined by normalizing to ?-actin mRNA levels (Fig. 2). CRH (10 μM) exerted no observable effect on levels of BDNF mRNA after 24 h (data not shown) but significantly increased levels of BDNF mRNA expression to 434 ± 154% compared with untreated control after 48 h (100 ± 22.85%; P < 0.05 vs. vehicle control, n = 3; Fig. 2). This increase was significantly inhibited with the additional presence of H89 (184 ± 134%; P < 0.05 vs. CRH, n = 3; Fig. 2) and was almost abolished when cells were cotreated with CRH in combination with either the Ca2+ chelator BAPTA-AM or the CRHR1 antagonist R121919 (BAPTA-AM, 69 ± 24%; R121919, 60 ± 1%; P < 0.05 vs. CRH, n = 3; Fig. 2).
FIG. 2. CRH induces BDNF expression in cerebellar granule neurons in a calcium- and PKA-dependent manner. Semiquantitative RT-PCR for BDNF using ?-actin as an internal standard is shown (one representative gel is shown). Neurons were treated with CRH (10–8 M) alone or in combination with H89 (10–5 M), BAPTA-AM (BAPTA; 10–6 M), or R121919 (100 ng/ml) for 48 h. Results were calculated as ratios of OD of the BDNF band vs. the ?-actin band and expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. untreated control; #, P < 0.05 vs. CRH (n = 4).
WIN-55,212-2 inhibits depolarization-induced BDNF expression in cerebellar cultures
The activation of calcium-dependent pathways has been well documented in controlling BDNF expression, and thus, we examined the putative effects of CB1 activation on calcium-dependent, depolarization-induced BDNF expression (9). BDNF ELISAs were carried out with protein extracts from mouse cerebellar neurons initiated from CB1-deficient mice and wild-type littermates. We observed that depolarization-induced (KCl, 35 mM; 48 h) BDNF expression was significantly inhibited by application of the CB1 agonist WIN-55,212-2 (10–6 M) in wild-type mice (CB1+/+: KCl, 3051 ± 732.5%; KCl + WIN-55,212-2, 400.7 ± 87.6%; P < 0.05 vs. vehicle control, 100 ± 30.99%; P < 0.01 vs. KCl; n = 3; Fig. 3). However, inhibitory effects of WIN-55,212-2 (10–6 M) were absent in CB1-deficient mice (CB1–/–: KCl, 2960 ± 521.8%; KCl + WIN-55,212-2, 2406 ± 932.8%; P < 0.05 vs. vehicle control, 100 ± 20.38%; n = 3; Fig. 3), thereby implying that inhibitory effects on depolarization-induced BDNF expression are mediated by CB1 receptor activation.
FIG. 3. WIN-55,212-2 inhibits calcium-induced, activity-dependent BDNF expression in a CB1-dependent manner. BDNF ELISA from cerebellar cultures initiated either from CB1-deficient pups (CB1–/–) and wild-type littermates (CB1+/+). Cultures were treated with KCl (35 mM) alone or in combination with WIN-55,212-2 (WIN; 10–6 M) for 48 h. Samples were measured in duplicates, and data were expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. vehicle control (veh); ##, P < 0.01 vs. KCl (n = 3).
CRH-mediated increase of BDNF expression is inhibited by CB1 activation
To analyze the effects of CB1 activation on CRH-induced BDNF expression, we carried out semiquantitative RT-PCR. Total RNA extracted from cultures of rat cerebellar neurons was used treated with CRH (10–8 M), WIN-55,212-2 (10–6 M), or a combination of both. Relative expression was determined by normalizing to the levels of the housekeeping gene HPRT mRNA (Fig. 4A). Treatment of neurons with CRH alone resulted in an increase in BDNF mRNA transcripts to 200 ± 40.76% (P < 0.05 vs. vehicle control, 100 ± 13.25%; n = 4; Fig. 4A) after 48 h but not after 24 h (data not shown). Simultaneous treatment with WIN-55,212-2 significantly reduced the stimulatory effect of CRH (CRH + WIN-55,212-2, 128.3 ± 10.61%; P < 0.05 vs. CRH; n = 4). WIN-55,212-2 alone had no observable effects on BDNF expression compared with vehicle control (Fig. 4A). Using ELISA, it was observed that BDNF protein levels were unchanged 24 h after stimulation with CRH (data not shown), but significantly increased BDNF protein levels were observed after 48 h (160.1 ± 63.45%; P < 0.05 vs. vehicle control, 100 ± 34.45%; n = 3; Fig. 4B). This increase was inhibited by the addition of WIN-55,212-2 (CRH + WIN-55,212-2, 86.86 ± 16.17%; P < 0.05 vs. CRH; n = 3; Fig. 4B). The CB1 antagonist SR141716A (5 μM) blocked the inhibitory effect of WIN-55,212-2 on CRH-induced BDNF stimulation (CRH + WIN-55,212-2 + SR141716A, 94.54 ± 27.31%; P < 0.05 vs. CRH; n = 3; Fig. 4B). Application of WIN-55,212-2 alone did not show any significant changes of BDNF protein levels compared with vehicle control (Fig. 4B).
FIG. 4. Effects of WIN-55,212-2 on CRH-induced BDNF expression in cerebellar granule neurons. A, Semiquantitative RT-PCR for BDNF, using HPRT as an internal standard, is shown (one representative gel is shown). Neurons were treated with CRH (10–8 M) and/or WIN-55,212-2 (WIN; 10–6 M) for 48 h. Results were calculated as ratios of OD of the BDNF band vs. the HPRT band and expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. vehicle control (veh); #, P < 0.05 vs. CRH (n = 4). B, BDNF ELISA from cerebellar cultures initiated from rat pups. Neurons were treated with CRH (10–8 M) alone, WIN-55,212-2 (WIN; 10–6 M) alone, CRH and WIN-55,212-2, or CRH, WIN-55,212-2, and SR141716A (SR1) for 48 h. Samples were measured in duplicates, and data were expressed as the mean ± SEM of the percentage of basal BDNF levels (considered as 100%). *, P < 0.05 vs. vehicle control (veh); #, P < 0.05 vs. CRH (n = 3).
Inhibition of CRH-mediated signaling by CB1 activation
CRH-induced activation of CRHR1 leads to the production of cAMP through stimulation of AC by Gs. Conversely, CB1 has been reported to inhibit AC through a Gi/o-mediated mechanism. A possible site of interaction between the CRH and cannabinoid systems may center upon cAMP production. Therefore, we analyzed the effects of the CB1 agonist WIN-55,212-2 on CRH-induced cAMP accumulation in cultures from postnatal cerebellar granular neurons. Although the application of CRH (10–8 M) for 10 min induced a significant increase in intracellular cAMP levels to 259.6 ± 17.04% (P < 0.05 vs. vehicle control, 100 ± 34.58%; n = 6; Fig. 5A), simultaneous incubation for 10 min with the CB1 agonist WIN-55,212-2 (10–6 M) reduced CRH-mediated cAMP production, although it did not reach significant differences compared with CRH alone (CRH + WIN-55,212-2, 193.6 ± 17.92%; CRH, 259.6 ± 17.04%; P < 0.05 vs. CRH, n = 6; Fig. 5A). WIN-55,212-2 alone had no effect on intracellular cAMP levels (Fig. 5A).
FIG. 5. Modulation of CRH-induced signaling by WIN-55,212-2 in cerebellar granule neurons. A, cAMP accumulation assays with neurons treated for 10 min with CRH (10–8 M) and/or WIN-55,212-2 (WIN; 10–6 M) are shown. Samples were measured in triplicates, and data are expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. vehicle control (veh; n = 6). B and C, Western blot analyzing the phosphorylation status of CREB (two representative blots are shown). Phosphorylated levels of protein were normalized to total unphosphorylated levels and depicted as percent increase ± SEM of vehicle control (considered as 100%). B, Neurons were treated for 30 min with CRH (10–8 M) and/or WIN-55,212-2 (WIN; 10–6 M). *, P < 0.05 vs. vehicle control (veh; n = 3); #, P < 0.05 vs. CRH (n = 3). C, Analysis of the concentration-dependent effects of WIN-55,212-2 (WIN; 10–6–10–9 M) on CREB phosphorylation induced by CRH. *, P < 0.05 vs. vehicle control (veh; n = 3); #, P < 0.05 vs. CRH (n = 3).
The effect of CB1 activation on CREB phosphorylation was also investigated. CREB, a transcription factor that controls BDNF expression, is a downstream target of cAMP (34). Increases in intracellular cAMP concentrations lead to the activation of PKA, which in turn translocates to the nucleus and promotes the phosphorylation of CREB. CREB-dependent gene transcription is initially controlled by phosphorylation and subsequent activation at serine 133. Modulatory effects of CB1 activation on CRH-mediated signaling were monitored by Western blot analysis carried out using phospho-specific antibodies directed against the activated form of CREB, phospho-CREB (pCREB). Treatment of cerebellar neurons with CRH (10–8 M) for 30 min led to an increase of pCREB levels to 469 ± 153% (P < 0.05 vs. vehicle control, 100 ± 28.86%; n = 3; Fig. 5B). However, cotreatment of cell cultures with WIN-55,212-2 (10–6 M) for 30 min inhibited CRH-mediated CREB phosphorylation (128 ± 27% of vehicle control; P < 0.05 vs. CRH; n = 3; Fig. 5B). Furthermore, we carried out a dose-response curve of the inhibitory effect of WIN-55,212-2 (10–6-10–9 M) on CRH-mediated CREB phosphorylation (Fig. 5C). Significant inhibitory effects were observed at concentrations of WIN-55,212-2 ranging from 10–6-10–8 M (CRH vs. CRH/WIN-55,212-2; decrease of pCREB from 516 ± 202% to 134 ± 22.04%, 126 ± 7.4%, and 214 ± 46%, for 10–6, 10–7, and 10–8 M of WIN-55,212-2, respectively; P < 0.05 in each case).
Discussion
Our results demonstrate a functional interaction between the CRH and the cannabinoid systems that occurs at the level of second messenger signaling and that affects the regulation of BDNF gene expression. In situ hybridization experiments on sections from mouse cerebellum showed high levels of coexpression in granule neurons, which is a necessary prerequisite for any direct interaction. Moreover, coexpression of both receptors was also detected in numerous forebrain regions (35). CRH was demonstrated to induce an increase in cAMP and pCREB levels, as well as elevations in BDNF transcripts and protein in a calcium-dependent manner. These effects were inhibited by activation of CB1 receptors. Moreover, CB1 receptor activation inhibited depolarization-induced increases of BDNF protein. Therefore, this study reveals an important interaction between the CRH and cannabinoid systems in the modulation of BDNF expression by influencing cAMP and Ca2+ signal transduction pathways (summarized in Fig. 6).
FIG. 6. A schematic representation showing the intracellular pathways that may mediate the interaction between CRHR1 and CB1, leading to the regulation of BDNF expression. Stimulation of CRHR1 with CRH activates AC, and subsequent cAMP production leads to the activation of PKA. Activated PKA phosphorylates CREB, which initiates the expression of the BDNF. Moreover, PKA is known to phosphorylate Ca2+ channels, which would lead to an increased influx of Ca2+ and thus activation of BDNF expression. Depolarization of the cells with KCl (35 mM) also leads to increased BDNF expression. Concurrent stimulation of CB1 with its agonist WIN-55,212-2 (WIN) inhibits CRH- and depolarization-induced BDNF expression through its negative effect on the cAMP signaling cascade and its ability to inhibit the activity of Ca2+ channels.
CRHR1 is a seven-transmembrane domain receptor linked to AC through Gs protein activation. Subsequent cAMP production leads to the activation of PKA, a mechanism that mediates the majority of CRH-mediated effects. Classically, CRH-mediated release of ACTH is PKA dependent (5), and recently, the neuroprotective effects associated with CRHR1 activation were shown to be PKA dependent (36). It is well established that PKA activation leads to the phosphorylation and, hence, activation of CREB. Additionally, PKA is known to phosphorylate L-type Ca2+ channels (37). In a manner similar to other seven-transmembrane receptors, CRHRs may also activate this channel type in neurons, resulting in an increased Ca2+ influx (38). L-type Ca2+ channels have indeed been demonstrated to be involved during the regulation of CRH-mediated ACTH release in corticotrope cells (32).
The regulation of BDNF expression by CREB has been studied extensively, and a majority of reports demonstrated a Ca2+-dependent mechanism of CREB phosphorylation and induction of BDNF expression (33). In cortical neurons, a cooperation between forskolin-induced PKA activation and Ca2+ influx triggers phosphorylation of CREB, followed by binding to the Ca2+-dependent response element within the BDNF gene (34). Moreover, it was shown that forskolin mediates increases in BDNF expression in raphe neurons in a PKA-dependent manner (8). Our results demonstrate that CRH enhances BDNF transcription in a Ca2+-, PKA-, and CRHR1-dependent manner in cerebellar neurons. Taken together, these data indicate that CRH is able to induce BDNF expression by activation of the cAMP/PKA signaling cascade and by an increase of intracellular Ca2+ concentrations, possibly due to an activation of L-type Ca2+ channels.
CB1 receptors also seem to participate in the Ca2+-dependent regulation of BDNF expression because we showed that depolarization-induced increases of BDNF protein were inhibited by the CB1 receptor agonist WIN-55,212-2 in neuronal cultures from wild-type mice but not in cultures from CB1-deficient animals. Cannabinoids were found to inhibit N- and P/Q-type voltage-dependent calcium currents in several cell lines (39, 40) as well as in primary cultures of cerebellar granule neurons (41) via pertussis toxin-sensitive G proteins. In addition, another study using cerebral artery smooth muscle cells of the cat also showed an inhibition of L-type Ca2+ channels upon activation of CB1 with WIN-55,212-2 and the endocannabinoid anandamide (42). On the other hand, the cannabinoid receptor agonist desacetyl levonantradol is able to increase Ca2+ influx into the neuroblastoma cell line N18TG2 at nanomolar concentrations (43). This effect is mediated by Gs GTP-binding proteins (44). Because CREB phosphorylation and activation of BDNF transcription are preferentially driven by calcium influx through L-type Ca2+ channels, whereas they are poorly induced by calcium entering through N-methyl-D-aspartate receptors and non-L-type Ca2+ channels (9, 45), the possible role of L-type Ca2+ channels in CB1-mediated inhibition of CRH-induced effects remains to be further investigated in detail.
A main feature of CB1 effects is the inhibition of AC via Gi/o. This has been observed in a number of cell types, including neuroblastoma cells (46), in CB1-transfected cell lines (47), and in rat cerebellar granule cells (48). Cannabinoid-induced inhibition of AC results in the attenuation of PKA activity and a decrease in binding of transcription factors to cAMP response elements present in target genes (49). Additionally, independent of its activation state, CB1 is able to sequester G, G?, and G proteins required by other receptors linked with pertussis toxin-sensitive Gi/o proteins (50). CRHRs are highly promiscuous because they can activate many different types of G proteins. In the rat cerebral cortex, CRHRs can activate Gs, Gi, Gq, and Gz (51). Therefore, both inhibition of CRH-mediated cAMP augmentation and sequestration of G proteins required by the CRHR1 may be involved in the inhibitory action of CB1 on CRH-mediated signaling and induction of BDNF expression.
The inhibition of CRH-induced intracellular signaling by endocannabinoids has important implications with respect to the process of glucocorticoid inhibition. The first report of rapid inhibition of CRH neurons in the hypothalamic paraventricular nucleus by corticosteroids has recently identified endocannabinoids as being mediators of nongenomic corticosteroid actions (21). The proposed mechanism involves glucocorticoid binding to a G protein-coupled glucocorticoid receptor and subsequent activation of intracellular signaling pathways that lead to endocannabinoid synthesis and release. The released endocannabinoids act as retrograde messengers that bind to CB1 receptors presynaptically, which leads to an inhibition of CRH release. Thus, in light of the data presented here, we suggest that corticosteroids may additionally inhibit CRH-mediated activation of intracellular signaling pathways and gene expression postsynaptically through similar endocannabinoid-dependent mechanisms.
Endocannabinoids and 9-tetrahydrocannabinol have been demonstrated to activate ERK in hippocampal slices and BDNF expression in vivo (52). The effects of cannabinoids were dependent on N-methyl-D-aspartate receptor activation in vivo but not in hippocampal slices, suggesting that multiple pathways lead to the initiation of CB1-mediated signaling pathways. In the present study, CB1 activation with the agonist WIN-55,212-2 did not result in increases in BDNF expression in cultured cerebellar granule neurons. This may be due to region- and cell-specific coupling of signaling pathways, as has been demonstrated in the case of CRH (36), or interactions with other systems.
Because we were able to show CRH-induced BDNF expression after 48 h, physiological long-term changes, such as synaptic plasticity, might be the consequence of this altered gene expression. Deprivation of BDNF leads to an impairment of hippocampal-dependent long-term potentiation (LTP), a form of synaptic plasticity, suggesting that BDNF is essential for certain forms of learning and memory (53). Because a brain region-specific signaling profile of CRH was identified, indicating that CRH promotes common signaling pathways in cerebellar and hippocampal but not cortical neurons (36), we suggest that CRH may affect similar processes in the cerebellum. In contrast to CRH, learning and memory impairments are among the most commonly reported behavioral effects of exogenous cannabinoids (for review, see Ref.54). On the other hand, the temporally and spatially restricted release of endocannabinoids facilitates the induction of LTP in the hippocampus in single neurons (55). Because we activated CB1 in all cells by exogenously applied WIN-55,212-2 and CB1 receptors are located on excitatory glutamatergic synapses of granule neurons (56) in our model system, we would rather suggest that there are negative effects on LTP. Moreover, previous work has indicated that induction of cerebellar LTP, which is involved in certain forms of motor learning, including associative eyeblink conditioning and adaptation of the vestibulo-ocular reflex, requires presynaptic Ca2+ influx (57), cAMP production (58), and activation of PKA (59), which are all negatively influenced by CB1 activation. Thus, exogenously applied cannabinoids might mediate their negative effects on learning and memory through dampening the signaling pathway of other neurotransmitter systems that improve cognitive functions and, therefore, regulating the expression of downstream target genes.
Taken together, our study highlights an important role for CRH in regulating BDNF expression. The function of CB1 in inhibiting the CRH signaling cascade is consistent with the role that CB1 ligands generally play. This encompasses the mechanisms by which cannabinoids negatively regulate the cAMP-PKA pathway (46, 49) and also included effects on Ca2+-dependent signaling pathways, which are crucial in the modulation of the expression of the neurotrophins and, in particular, BDNF. The general pharmacological stimulation of CB1 receptors clearly has a negative effect on plasticity (49, 60), which might be mediated by inhibiting signaling pathways, leading to expression changes of target genes. Therefore, further studies should concentrate on the molecular and intracellular interactions of CRH and the endocannabinoid system, not only in terms of stress and glucocorticoid-mediated regulation of the HPA axis, but also in terms of the interactions of these two systems in diverse memory processes with particular attention to the regulation of expression and neuronal release of neurotrophins.
Acknowledgments
We thank Dr. G. Marsicano for statistical evaluation and fruitful discussions and for providing the CB1-deficient mouse line, J. Zschocke and Dr. A. Clement for critical reading of the manuscript, B. W?lfel for excellent support in animal work, and F. Ohl for supplying R121919. CRHR1 plasmid was a kind gift from Dr. W. Wurst.
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Address all correspondence and requests for reprints to: Dr. Christian Behl, Department of Pathobiochemistry, Johannes Gutenberg University Mainz Medical School, 55099 Mainz, Germany. E-mail: cbehl@uni-mainz.de.
Abstract
CRH receptor (CRHR) 1 and the cannabinoid receptor 1 (CB1) are both G protein-coupled receptors. Activation of CRHR1 leads to increases in cAMP production and phosphorylation of the transcription factor cAMP response element-binding protein (CREB). In contrast, CB1 is negatively coupled to the cAMP signaling cascade. In this study, we analyzed a putative interaction between these two systems focusing on the regulation of the expression of brain-derived neurotrophic factor (BDNF), a CREB-regulated gene. In situ hybridization revealed coexpression of CRHR1 and CB1 receptors in the granular layer of the cerebellum. Therefore, we analyzed the effects of CRH and the CB1 agonist WIN-55,212-2 on BDNF expression in primary cerebellar neurons from rats and mice. We observed that application of CRH for 48 h led to an increase in BDNF mRNA and protein levels. This effect was inhibited by WIN-55,212-2. At the level of intracellular signaling, short-term application of WIN-55,212-2 inhibited CRH-induced cAMP accumulation and CREB phosphorylation. Pharmacological analysis demonstrated that the CRHR1 antagonist R121919, the protein kinase A inhibitor H89, and the calcium chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid acetoxymethyl ester inhibited CRH-mediated BDNF expression. Moreover, depolarization-induced BDNF synthesis was also inhibited by long-term application of WIN-55,212-2 in wild-type mice but not in CB1-deficient mice. Thus, these data highlight an interaction between the CRH and the cannabinoid system in the regulation of BDNF expression by influencing cAMP and Ca2+ signaling pathways.
Introduction
CRH, A MAJOR mediator of the stress response in the central nervous system (CNS) (1) also affects other central processes, such as learning and memory, synaptic plasticity, and neuroprotection (2, 3, 4). These effects are mediated by CRH binding and activation of two distinct G protein-coupled receptors, CRH receptor (CRHR) 1 and CRHR2, which are found throughout the CNS and the periphery (5). CRH has a higher affinity for CRHR1 than for CRHR2, and in the brain, CRHR1 is expressed at high levels in the hippocampus, cortex, and cerebellum (6). CRH binding to CRHR1 typically activates adenylate cyclase (AC), which leads to increased intracellular concentrations of cAMP and activation of protein kinase A (PKA) (5). However, downstream target genes of CRH have not yet been thoroughly investigated; previous studies have focused on genes involved in the stress response and ACTH release, such as proopiomelanocortin (7). One putative target is the brain-derived neurotrophic factor (BDNF), the expression of which is controlled by cAMP-elevating agents in neurons (8). In addition to its role as a classical target-derived growth factor during neuronal development, BDNF is an essential autocrine factor, released and acting locally after neuronal depolarization (9).
Cannabinoids also exert most of their effects through activation of G protein-coupled receptors, which include the inhibition of AC through Gi/o proteins (10). To date, two cannabinoid receptors have been identified; the cannabinoid receptor type 1 (CB1) is widely expressed throughout the CNS (11), whereas the cannabinoid receptor type 2 is mainly present in immune cells (12). Among other regions, CB1 is localized densely in the cerebellum of the postnatal and adult brain (13, 14).
A striking feature of the cannabinoid system is its involvement in the modulation of various neurotransmitter systems, mainly by decreasing neurotransmitter release (15). Despite the recent progress in understanding the actions of endocannabinoids on synaptic transmission (16), the signal transduction pathways regulated by Gi/o-coupled CB1 in the cerebellum are poorly characterized. Most of the data were obtained in nonneuronal cell lines, in which stimulation of CB1 activates the ERK subtype of MAPK (17) eventually inducing the expression of the immediate-early genes coding for c-fos and Zif268 (18). Although there is evidence that activation of CB1 by exogenous ligands (9-tetrahydrocannabinol and WIN-55,212-2) regulates the release of ACTH via the secretion of CRH within the hypothalamic-pituitary-adrenal (HPA) axis (19, 20), signaling interactions between the corresponding receptors and subsequent effects on downstream transcriptional targets have not been investigated in detail. Intriguingly, a rapid component of glucocorticoid-negative feedback, the process that controls the stress response by affecting the HPA axis through the inhibition and chronic reduction of expression of CRH in the hypothalamus, has been recently reported to be mediated in a cannabinoid/CB1-dependent manner (21).
In the present study, we investigated other putative modes of interaction between the cannabinoid system and the CRH system regarding intracellular signaling mechanisms. As a model system, we used primary cultures of cerebellar granule neurons where both receptors are highly expressed. Cerebellar granule cells constitute the largest homogeneous neuronal population of mammalian brain. Due to their postnatal generation and the feasibility of well-characterized primary in vitro cultures, cerebellar granule neurons have acquired a special position in neuroscience as one of the most reliable models for the study of neural development, function, and pathology. We examined the effects of CRH and the CB1 agonist WIN-55,212-2 on BDNF expression, the transcription of which is reported to be dependent on cAMP response element-binding protein (CREB) phosphorylation and neuronal activity and might, therefore, represent a downstream target.
Materials and Methods
Drugs
3-Isobutyl-1-methylxanthine (IBMX; Sigma, Deisenhofen, Germany), WIN-55,212-2 (Tocris, Cologne, Germany), rat/human CRH (Calbiochem, Schwalbach, Germany), 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid acetoxymethyl ester (BAPTA-AM) (Calbiochem), H89 (Calbiochem), and R121919 (kind gift from Dr. F. Ohl, Max Planck Institute of Psychiatry, Munich, Germany) were prepared as concentrated stock solutions in PBS (IBMX and H89, both 10 mM), 100% dimethylsulfoxide (DMSO; WIN-55,212-2, 10 mM, and BAPTA-AM, 1 mM), and 2% acetic acid (CRH, 10 mM) or citrate buffer (R121919, 100 mg/ml).
Animals and tissue preparation
Sprague Dawley rats, C57BL/6N mice (both from Charles River, Sulzfeld, Germany), CB1-deficient mice, and wild-type littermates (22) were housed with a 12-h light, 12-h dark cycle and allowed to access food and water ad libitum. All experimental protocols were approved by the Ethical Committee on Animal Care and Use of the Governments of Bavaria and Rheinland-Palatinate, Germany.
Adult mice (3–5 months old) were killed by cervical dislocation for in situ hybridization experiments, and newborn rats or mice were decapitated for cerebellar primary cultures. Brains were snap frozen on dry ice immediately after isolation and stored at –80 C until sectioning. Brains were mounted on Tissue Tek (Polysciences, Warrington, PA), and 18-μM-thick coronal consecutive sections were cut from the cerebellum on a cryostat (Microtome HM 560; Microm, Braunschweig, Germany). Sections were mounted onto frozen SuperFrost/plus slides (Fisher Scientific, Schwerte, Germany), dried on a 42 C warming plate, and stored at –20 C until used.
Synthesis of probes
35S-labeled and digoxigenin (DIG)-labeled riboprobes were used. CRHR1 plasmid was a kind gift from Dr. W. Wurst (Max Planck Institute of Psychiatry) and was used as described previously (23). The CB1 probe was used as described in Marsicano and Lutz (24). In vitro transcription was carried out for 35S-labeled and DIG-labeled riboprobes, as previously described (25). Using these probes for in situ hybridization experiments, sense controls did not give any detectable signals, and antisense probes gave expression patterns identical to those already published in rat or mouse (data not shown).
In situ hybridization
Single in situ hybridization with 35S-labeled riboprobes for CB1 and CRHR1 was carried out as previously described (24), and double in situ hybridization was carried out as previously described (25). The TSA Biotin System (NEN Life Science Products, Boston, MA) was used for detection of the DIG-labeled CB1 probe, and the chromogenic reaction was carried out with the Vector Red kit (Vector Laboratories, Gruenberg, Germany). Slides were dipped in photographic emulsion (NTB-2 from Kodak, Rochester, NY; diluted 1:1 in distilled H2O) for detection of the 35S-labeled CRHR1. After 4 wk of exposure at 4 C, slides were developed (D-19; Kodak) and fixed (Kodak fixer).
Cell culture
Primary rat or mouse cerebellar granular cultures were prepared as described previously with modifications (26). Cerebellar tissue from 3-d-old animals was dissected, and tissue pieces were incubated for 20 min in Ca2+-free, Mg2+-free Dulbecco’s PBS (Invitrogen, Karlsruhe, Germany) containing 0.1% trypsin and 0.02% EDTA (pH 8.0) at room temperature. Cells were transferred to Ca2+-free, Mg2+-free Hank’s balanced salt solution (Invitrogen) supplemented with 10% fetal calf serum (Invitrogen) and dissociated gently by pipetting up and down. Undissociated pieces were filtered through a 50-μM pore-sized Nybolt mesh (Eckardt, Waldkirch, Germany), and cells were centrifuged at 200 x g for 4 min. The pellet was resuspended in MEM (Invitrogen) supplemented with 10% horse serum (Invitrogen), and the number of viable cells was counted by trypan blue exclusion. Cells were seeded (150,000 cells/cm2) into poly-L-ornithine-coated (0.1 mg/ml; molecular mass, 100–200 kDa; Sigma) 6- or 24-well plates with MEM (Invitrogen) supplemented with 10% horse serum in the case of rat cultures or MEM supplemented with 20% fetal calf serum in the case of mouse cultures. Culture medium was changed to serum-free N2-supplemented MEM/F12 (Invitrogen) medium after 24 h for rat cultures and Neurobasal medium supplemented with B27 (both from Invitrogen) for mouse cultures. Cells were used for experiments after another 24 h.
cAMP accumulation assay
The cAMP assay was performed as described previously (27) with slight modifications. Primary cultures were maintained in 24-well plates, and IBMX was added to cultures 5 min before addition of CRH (10–8 M) and/or WIN-55,212-2 (10–6 M). Cells were incubated for 10 min with the drugs, and reactions were terminated by aspiration of the medium and addition of 1 ml ice-cold 6% trichloroacetic acid, followed by incubation overnight at 4 C. Two percent acetic acid alone and 0.2% DMSO used as vehicle controls had no effect on cAMP accumulation. The extracts were treated twice with 4 ml diethylether to remove the trichloroacetic acid, dried overnight in a lyophilisator, and reconstituted in serum-free culture medium. Intracellular cAMP levels were measured with a competitive protein binding assay (NEN Life Science Products). Data are expressed as percentage of basal cAMP levels. Extracts from two wells were pooled, and samples were measured in triplicates.
Immunoblot analysis
Cells plated in six-well dishes were lysed in 200 μl lysis buffer (60 mM Tris/HCl, pH 8.0; 2% sodium dodecyl sulfate, 10% sucrose, 5 μg/ml aprotinin, and 10 mM phenylmethylsulfonyl fluoride) per well. Lysates were briefly sonicated and stored at –80 C. Protein content was measured using the BCA assay (Pierce, Bonn, Germany). Samples were diluted 1:1 in 2x sample buffer (250 mM Tris-HCl, pH 6.8; 4% sodium dodecyl sulfate, 10% glycerol, 2% ?-mercaptoethanol, and 0.0006% bromophenol blue) and denatured at 95 C for 5 min. Proteins (10 μg/lane) were separated by SDS-PAGE (10% resolving gel) and transferred onto a nitrocellulose membrane (Schleicher and Schuell, Dassel, Germany) by semidry transfer (Bio-Rad, Munich, Germany). Blots were blocked for 30 min at room temperature with blocking buffer (5% fat-free milk powder, 0.05% Tween 20, 20 mM Tris/HCl, pH 7.6; and 150 mM NaCl) and incubated with either anti-CREB (1:500; Calbiochem) or anti-pCREB (phosphorylated at serine 133; Upstate Biotechnologies, Lake Placid, NY) primary antibodies in blocking buffer overnight at 4 C. After incubation with a horseradish peroxidase-coupled secondary antibody for 1.5 h (1:5000; Dianova, Hamburg, Germany) at room temperature, bands were visualized using the ECL detection system (Pharmacia, Freiburg, Germany). ODs of bands were calculated with Scion software (Scion Corp., Frederick, MA). Phosphorylated protein levels were normalized to total unphosphorylated levels.
Semiquantitative RT-PCR
Total RNA was isolated from cultures with peqGOLD RNAPure (Peqlab, Erlangen, Germany) according to the manufacturer’s instructions. Residual genomic DNA was removed with RNase-free DNase I. RNA (5 μg) was used for Superscript II (Life Technologies, Karlsruhe, Germany) reverse transcriptase-mediated synthesis of oligo(dT)12–18-primed (Roche, Mannheim, Germany) cDNA. PCR was carried out as follows: 94 C for 1 min; 55 C for hypoxanthinguanine phosphoribosyl transferase (HPRT) and ?-actin or 63 C for BDNF for 1 min; and 72 C for 1 min, with a 10-min extension at 72 C during the last cycle. PCR was carried out with 28 and 32 cycles for HPRT/?-actin and BDNF, respectively. Primer sequences included the following: BDNF (28); HPRT: sense 5'-CCTGCTGGATTACATTAAAGCACTG-3'; antisense 5'-GTC AAG GGC ATA TCC AAC AAA C-3'; and ?-actin: sense 5'-CTA CAA TGA GCT GCG TGT GGC-3'; antisense 5'-CAG GTC CAG ACG CAG GAT GGC-3'. PCR products of 297, 351, and 275 bp for BDNF, HPRT, and ?-actin, respectively, were amplified. Negative RNA controls without reverse transcriptase ensured a lack of genomic DNA contamination. ODs of PCR bands were measured with the Kodak-1D software. Results were calculated as ratios of OD of the BDNF vs. HPRT or ?-actin bands, respectively. To ensure that the PCR products fell within the linear range, cycle dependency was carried out for BDNF, HPRT and ?-actin. PCR samples were run on agarose gels, and ODs were measured using Scion Image software (Scion Corp.) and plotted on graphs. In each case, the correlation coefficient was r2 0.97 (data not shown).
ELISA for BDNF expression
The Emax immunoassay system (Promega, Mannheim, Germany) was used to quantify the levels of BDNF protein in primary neuronal cultures. DMSO (0.2%) vehicle control had no effect on BDNF expression. Lysis of cells, determination of protein content, and ELISA procedure were carried out as described in detail elsewhere (29). Samples from three pooled wells were measured in duplicates.
Statistical analysis
Data were analyzed by one-way ANOVA using GraphPad (GraphPad Prism, San Diego, CA) or SigmaStat (Systat Software, Point Richmond, CA) software. Significance between groups was further analyzed using the post hoc Tukey test, and data were always depicted with SEM. P < 0.05 was considered statistically significant.
Results
Colocalization of CB1 and CRHR1 in the mouse cerebellum
Both CB1 and CRHR1 receptors have been described to be expressed in the postnatal and adult cerebellum. In adult mouse and rat cerebellum, CRHR1 receptor mRNA and protein was described to be present throughout the cerebellum in Purkinje cells, the granule cell layer, and within all four of the cerebellar nuclei (30). Binding studies in early postnatal mice (postnatal d 3) showed CRHR1 distribution throughout the cerebellum. At postnatal d 10, the adult pattern of distribution and level of labeling begins to emerge (31). Similarly, CB1 was detected in the postnatal rat cerebellum by binding studies, in situ hybridization and immunohistochemistry showed an increasing signal density in adulthood, where it is mainly expressed throughout the granule and molecular layers (13, 14). However, the locations of both CB1 and CRHR1 within the same cells must be demonstrated to put forward a putative interaction between these two receptor systems.
In Fig. 1, we show two sections of the cerebellum that were hybridized with 35S-labeled riboprobes for either CB1 (Fig. 1A) or CRHR1 (Fig. 1B), indicating the high expression levels of both receptors in this layer. To show that both receptors are indeed present in the same cells, we performed double in situ hybridization by combining a DIG-labeled riboprobe for CB1 with a 35S-labeled riboprobe for CRHR1. Due to the high density of cells in the granule layer, it was not feasible to count coexpressing cells at a single-cell resolution in this area. Because silver grains (expression of CRHR1) and red staining (expression of CB1) are uniformly distributed throughout all layers of granule cells, we estimated a 100% coexpression of CB1 and CRHR1 receptors (Fig. 1C).
FIG. 1. Bright- and dark-field micrographs of coronal cerebellar sections showing examples of coexpression of CB1 with CRHR1 as detected by single or double in situ hybridization, respectively. Expression of (A) CB1 and (B) CRHR1 in the cerebellum as detected with 35S-labeled riboprobes. Scale bars, 1 mm. C, Coexpression of CB1 (red staining) and CRHR1 (silver grains) in the granule layer. Scale bar, 50 μm. ML, Molecular layer; GL, granular layer.
CRH induces BDNF expression in cerebellar cultures through calcium- and PKA-dependent signaling pathways
The majority of the actions of CRH have been demonstrated to be mediated through the activation of AC and the subsequent elevation in intracellular cAMP levels (5). Moreover, CRH-mediated ACTH release in corticotrope cells has been shown to involve L-type Ca2+ channels (32). Elevations in intracellular cAMP or activation of Ca2+-dependent pathways result in the induction of BDNF expression (8, 33). Therefore, we examined the effects of CRHR1 activation on BDNF expression in cerebellar cultures. Semiquantitative RT-PCR analysis was carried out using BDNF-specific primers on cDNA derived from cells stimulated with CRH alone or in combination with the PKA inhibitor H89 (10 μM), the calcium chelator BAPTA-AM (1 μM), and the CRHR1 antagonist R121919 (100 ng/ml), respectively. Relative expression was determined by normalizing to ?-actin mRNA levels (Fig. 2). CRH (10 μM) exerted no observable effect on levels of BDNF mRNA after 24 h (data not shown) but significantly increased levels of BDNF mRNA expression to 434 ± 154% compared with untreated control after 48 h (100 ± 22.85%; P < 0.05 vs. vehicle control, n = 3; Fig. 2). This increase was significantly inhibited with the additional presence of H89 (184 ± 134%; P < 0.05 vs. CRH, n = 3; Fig. 2) and was almost abolished when cells were cotreated with CRH in combination with either the Ca2+ chelator BAPTA-AM or the CRHR1 antagonist R121919 (BAPTA-AM, 69 ± 24%; R121919, 60 ± 1%; P < 0.05 vs. CRH, n = 3; Fig. 2).
FIG. 2. CRH induces BDNF expression in cerebellar granule neurons in a calcium- and PKA-dependent manner. Semiquantitative RT-PCR for BDNF using ?-actin as an internal standard is shown (one representative gel is shown). Neurons were treated with CRH (10–8 M) alone or in combination with H89 (10–5 M), BAPTA-AM (BAPTA; 10–6 M), or R121919 (100 ng/ml) for 48 h. Results were calculated as ratios of OD of the BDNF band vs. the ?-actin band and expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. untreated control; #, P < 0.05 vs. CRH (n = 4).
WIN-55,212-2 inhibits depolarization-induced BDNF expression in cerebellar cultures
The activation of calcium-dependent pathways has been well documented in controlling BDNF expression, and thus, we examined the putative effects of CB1 activation on calcium-dependent, depolarization-induced BDNF expression (9). BDNF ELISAs were carried out with protein extracts from mouse cerebellar neurons initiated from CB1-deficient mice and wild-type littermates. We observed that depolarization-induced (KCl, 35 mM; 48 h) BDNF expression was significantly inhibited by application of the CB1 agonist WIN-55,212-2 (10–6 M) in wild-type mice (CB1+/+: KCl, 3051 ± 732.5%; KCl + WIN-55,212-2, 400.7 ± 87.6%; P < 0.05 vs. vehicle control, 100 ± 30.99%; P < 0.01 vs. KCl; n = 3; Fig. 3). However, inhibitory effects of WIN-55,212-2 (10–6 M) were absent in CB1-deficient mice (CB1–/–: KCl, 2960 ± 521.8%; KCl + WIN-55,212-2, 2406 ± 932.8%; P < 0.05 vs. vehicle control, 100 ± 20.38%; n = 3; Fig. 3), thereby implying that inhibitory effects on depolarization-induced BDNF expression are mediated by CB1 receptor activation.
FIG. 3. WIN-55,212-2 inhibits calcium-induced, activity-dependent BDNF expression in a CB1-dependent manner. BDNF ELISA from cerebellar cultures initiated either from CB1-deficient pups (CB1–/–) and wild-type littermates (CB1+/+). Cultures were treated with KCl (35 mM) alone or in combination with WIN-55,212-2 (WIN; 10–6 M) for 48 h. Samples were measured in duplicates, and data were expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. vehicle control (veh); ##, P < 0.01 vs. KCl (n = 3).
CRH-mediated increase of BDNF expression is inhibited by CB1 activation
To analyze the effects of CB1 activation on CRH-induced BDNF expression, we carried out semiquantitative RT-PCR. Total RNA extracted from cultures of rat cerebellar neurons was used treated with CRH (10–8 M), WIN-55,212-2 (10–6 M), or a combination of both. Relative expression was determined by normalizing to the levels of the housekeeping gene HPRT mRNA (Fig. 4A). Treatment of neurons with CRH alone resulted in an increase in BDNF mRNA transcripts to 200 ± 40.76% (P < 0.05 vs. vehicle control, 100 ± 13.25%; n = 4; Fig. 4A) after 48 h but not after 24 h (data not shown). Simultaneous treatment with WIN-55,212-2 significantly reduced the stimulatory effect of CRH (CRH + WIN-55,212-2, 128.3 ± 10.61%; P < 0.05 vs. CRH; n = 4). WIN-55,212-2 alone had no observable effects on BDNF expression compared with vehicle control (Fig. 4A). Using ELISA, it was observed that BDNF protein levels were unchanged 24 h after stimulation with CRH (data not shown), but significantly increased BDNF protein levels were observed after 48 h (160.1 ± 63.45%; P < 0.05 vs. vehicle control, 100 ± 34.45%; n = 3; Fig. 4B). This increase was inhibited by the addition of WIN-55,212-2 (CRH + WIN-55,212-2, 86.86 ± 16.17%; P < 0.05 vs. CRH; n = 3; Fig. 4B). The CB1 antagonist SR141716A (5 μM) blocked the inhibitory effect of WIN-55,212-2 on CRH-induced BDNF stimulation (CRH + WIN-55,212-2 + SR141716A, 94.54 ± 27.31%; P < 0.05 vs. CRH; n = 3; Fig. 4B). Application of WIN-55,212-2 alone did not show any significant changes of BDNF protein levels compared with vehicle control (Fig. 4B).
FIG. 4. Effects of WIN-55,212-2 on CRH-induced BDNF expression in cerebellar granule neurons. A, Semiquantitative RT-PCR for BDNF, using HPRT as an internal standard, is shown (one representative gel is shown). Neurons were treated with CRH (10–8 M) and/or WIN-55,212-2 (WIN; 10–6 M) for 48 h. Results were calculated as ratios of OD of the BDNF band vs. the HPRT band and expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. vehicle control (veh); #, P < 0.05 vs. CRH (n = 4). B, BDNF ELISA from cerebellar cultures initiated from rat pups. Neurons were treated with CRH (10–8 M) alone, WIN-55,212-2 (WIN; 10–6 M) alone, CRH and WIN-55,212-2, or CRH, WIN-55,212-2, and SR141716A (SR1) for 48 h. Samples were measured in duplicates, and data were expressed as the mean ± SEM of the percentage of basal BDNF levels (considered as 100%). *, P < 0.05 vs. vehicle control (veh); #, P < 0.05 vs. CRH (n = 3).
Inhibition of CRH-mediated signaling by CB1 activation
CRH-induced activation of CRHR1 leads to the production of cAMP through stimulation of AC by Gs. Conversely, CB1 has been reported to inhibit AC through a Gi/o-mediated mechanism. A possible site of interaction between the CRH and cannabinoid systems may center upon cAMP production. Therefore, we analyzed the effects of the CB1 agonist WIN-55,212-2 on CRH-induced cAMP accumulation in cultures from postnatal cerebellar granular neurons. Although the application of CRH (10–8 M) for 10 min induced a significant increase in intracellular cAMP levels to 259.6 ± 17.04% (P < 0.05 vs. vehicle control, 100 ± 34.58%; n = 6; Fig. 5A), simultaneous incubation for 10 min with the CB1 agonist WIN-55,212-2 (10–6 M) reduced CRH-mediated cAMP production, although it did not reach significant differences compared with CRH alone (CRH + WIN-55,212-2, 193.6 ± 17.92%; CRH, 259.6 ± 17.04%; P < 0.05 vs. CRH, n = 6; Fig. 5A). WIN-55,212-2 alone had no effect on intracellular cAMP levels (Fig. 5A).
FIG. 5. Modulation of CRH-induced signaling by WIN-55,212-2 in cerebellar granule neurons. A, cAMP accumulation assays with neurons treated for 10 min with CRH (10–8 M) and/or WIN-55,212-2 (WIN; 10–6 M) are shown. Samples were measured in triplicates, and data are expressed as the mean ± SEM of the percentage of vehicle control (considered as 100%). *, P < 0.05 vs. vehicle control (veh; n = 6). B and C, Western blot analyzing the phosphorylation status of CREB (two representative blots are shown). Phosphorylated levels of protein were normalized to total unphosphorylated levels and depicted as percent increase ± SEM of vehicle control (considered as 100%). B, Neurons were treated for 30 min with CRH (10–8 M) and/or WIN-55,212-2 (WIN; 10–6 M). *, P < 0.05 vs. vehicle control (veh; n = 3); #, P < 0.05 vs. CRH (n = 3). C, Analysis of the concentration-dependent effects of WIN-55,212-2 (WIN; 10–6–10–9 M) on CREB phosphorylation induced by CRH. *, P < 0.05 vs. vehicle control (veh; n = 3); #, P < 0.05 vs. CRH (n = 3).
The effect of CB1 activation on CREB phosphorylation was also investigated. CREB, a transcription factor that controls BDNF expression, is a downstream target of cAMP (34). Increases in intracellular cAMP concentrations lead to the activation of PKA, which in turn translocates to the nucleus and promotes the phosphorylation of CREB. CREB-dependent gene transcription is initially controlled by phosphorylation and subsequent activation at serine 133. Modulatory effects of CB1 activation on CRH-mediated signaling were monitored by Western blot analysis carried out using phospho-specific antibodies directed against the activated form of CREB, phospho-CREB (pCREB). Treatment of cerebellar neurons with CRH (10–8 M) for 30 min led to an increase of pCREB levels to 469 ± 153% (P < 0.05 vs. vehicle control, 100 ± 28.86%; n = 3; Fig. 5B). However, cotreatment of cell cultures with WIN-55,212-2 (10–6 M) for 30 min inhibited CRH-mediated CREB phosphorylation (128 ± 27% of vehicle control; P < 0.05 vs. CRH; n = 3; Fig. 5B). Furthermore, we carried out a dose-response curve of the inhibitory effect of WIN-55,212-2 (10–6-10–9 M) on CRH-mediated CREB phosphorylation (Fig. 5C). Significant inhibitory effects were observed at concentrations of WIN-55,212-2 ranging from 10–6-10–8 M (CRH vs. CRH/WIN-55,212-2; decrease of pCREB from 516 ± 202% to 134 ± 22.04%, 126 ± 7.4%, and 214 ± 46%, for 10–6, 10–7, and 10–8 M of WIN-55,212-2, respectively; P < 0.05 in each case).
Discussion
Our results demonstrate a functional interaction between the CRH and the cannabinoid systems that occurs at the level of second messenger signaling and that affects the regulation of BDNF gene expression. In situ hybridization experiments on sections from mouse cerebellum showed high levels of coexpression in granule neurons, which is a necessary prerequisite for any direct interaction. Moreover, coexpression of both receptors was also detected in numerous forebrain regions (35). CRH was demonstrated to induce an increase in cAMP and pCREB levels, as well as elevations in BDNF transcripts and protein in a calcium-dependent manner. These effects were inhibited by activation of CB1 receptors. Moreover, CB1 receptor activation inhibited depolarization-induced increases of BDNF protein. Therefore, this study reveals an important interaction between the CRH and cannabinoid systems in the modulation of BDNF expression by influencing cAMP and Ca2+ signal transduction pathways (summarized in Fig. 6).
FIG. 6. A schematic representation showing the intracellular pathways that may mediate the interaction between CRHR1 and CB1, leading to the regulation of BDNF expression. Stimulation of CRHR1 with CRH activates AC, and subsequent cAMP production leads to the activation of PKA. Activated PKA phosphorylates CREB, which initiates the expression of the BDNF. Moreover, PKA is known to phosphorylate Ca2+ channels, which would lead to an increased influx of Ca2+ and thus activation of BDNF expression. Depolarization of the cells with KCl (35 mM) also leads to increased BDNF expression. Concurrent stimulation of CB1 with its agonist WIN-55,212-2 (WIN) inhibits CRH- and depolarization-induced BDNF expression through its negative effect on the cAMP signaling cascade and its ability to inhibit the activity of Ca2+ channels.
CRHR1 is a seven-transmembrane domain receptor linked to AC through Gs protein activation. Subsequent cAMP production leads to the activation of PKA, a mechanism that mediates the majority of CRH-mediated effects. Classically, CRH-mediated release of ACTH is PKA dependent (5), and recently, the neuroprotective effects associated with CRHR1 activation were shown to be PKA dependent (36). It is well established that PKA activation leads to the phosphorylation and, hence, activation of CREB. Additionally, PKA is known to phosphorylate L-type Ca2+ channels (37). In a manner similar to other seven-transmembrane receptors, CRHRs may also activate this channel type in neurons, resulting in an increased Ca2+ influx (38). L-type Ca2+ channels have indeed been demonstrated to be involved during the regulation of CRH-mediated ACTH release in corticotrope cells (32).
The regulation of BDNF expression by CREB has been studied extensively, and a majority of reports demonstrated a Ca2+-dependent mechanism of CREB phosphorylation and induction of BDNF expression (33). In cortical neurons, a cooperation between forskolin-induced PKA activation and Ca2+ influx triggers phosphorylation of CREB, followed by binding to the Ca2+-dependent response element within the BDNF gene (34). Moreover, it was shown that forskolin mediates increases in BDNF expression in raphe neurons in a PKA-dependent manner (8). Our results demonstrate that CRH enhances BDNF transcription in a Ca2+-, PKA-, and CRHR1-dependent manner in cerebellar neurons. Taken together, these data indicate that CRH is able to induce BDNF expression by activation of the cAMP/PKA signaling cascade and by an increase of intracellular Ca2+ concentrations, possibly due to an activation of L-type Ca2+ channels.
CB1 receptors also seem to participate in the Ca2+-dependent regulation of BDNF expression because we showed that depolarization-induced increases of BDNF protein were inhibited by the CB1 receptor agonist WIN-55,212-2 in neuronal cultures from wild-type mice but not in cultures from CB1-deficient animals. Cannabinoids were found to inhibit N- and P/Q-type voltage-dependent calcium currents in several cell lines (39, 40) as well as in primary cultures of cerebellar granule neurons (41) via pertussis toxin-sensitive G proteins. In addition, another study using cerebral artery smooth muscle cells of the cat also showed an inhibition of L-type Ca2+ channels upon activation of CB1 with WIN-55,212-2 and the endocannabinoid anandamide (42). On the other hand, the cannabinoid receptor agonist desacetyl levonantradol is able to increase Ca2+ influx into the neuroblastoma cell line N18TG2 at nanomolar concentrations (43). This effect is mediated by Gs GTP-binding proteins (44). Because CREB phosphorylation and activation of BDNF transcription are preferentially driven by calcium influx through L-type Ca2+ channels, whereas they are poorly induced by calcium entering through N-methyl-D-aspartate receptors and non-L-type Ca2+ channels (9, 45), the possible role of L-type Ca2+ channels in CB1-mediated inhibition of CRH-induced effects remains to be further investigated in detail.
A main feature of CB1 effects is the inhibition of AC via Gi/o. This has been observed in a number of cell types, including neuroblastoma cells (46), in CB1-transfected cell lines (47), and in rat cerebellar granule cells (48). Cannabinoid-induced inhibition of AC results in the attenuation of PKA activity and a decrease in binding of transcription factors to cAMP response elements present in target genes (49). Additionally, independent of its activation state, CB1 is able to sequester G, G?, and G proteins required by other receptors linked with pertussis toxin-sensitive Gi/o proteins (50). CRHRs are highly promiscuous because they can activate many different types of G proteins. In the rat cerebral cortex, CRHRs can activate Gs, Gi, Gq, and Gz (51). Therefore, both inhibition of CRH-mediated cAMP augmentation and sequestration of G proteins required by the CRHR1 may be involved in the inhibitory action of CB1 on CRH-mediated signaling and induction of BDNF expression.
The inhibition of CRH-induced intracellular signaling by endocannabinoids has important implications with respect to the process of glucocorticoid inhibition. The first report of rapid inhibition of CRH neurons in the hypothalamic paraventricular nucleus by corticosteroids has recently identified endocannabinoids as being mediators of nongenomic corticosteroid actions (21). The proposed mechanism involves glucocorticoid binding to a G protein-coupled glucocorticoid receptor and subsequent activation of intracellular signaling pathways that lead to endocannabinoid synthesis and release. The released endocannabinoids act as retrograde messengers that bind to CB1 receptors presynaptically, which leads to an inhibition of CRH release. Thus, in light of the data presented here, we suggest that corticosteroids may additionally inhibit CRH-mediated activation of intracellular signaling pathways and gene expression postsynaptically through similar endocannabinoid-dependent mechanisms.
Endocannabinoids and 9-tetrahydrocannabinol have been demonstrated to activate ERK in hippocampal slices and BDNF expression in vivo (52). The effects of cannabinoids were dependent on N-methyl-D-aspartate receptor activation in vivo but not in hippocampal slices, suggesting that multiple pathways lead to the initiation of CB1-mediated signaling pathways. In the present study, CB1 activation with the agonist WIN-55,212-2 did not result in increases in BDNF expression in cultured cerebellar granule neurons. This may be due to region- and cell-specific coupling of signaling pathways, as has been demonstrated in the case of CRH (36), or interactions with other systems.
Because we were able to show CRH-induced BDNF expression after 48 h, physiological long-term changes, such as synaptic plasticity, might be the consequence of this altered gene expression. Deprivation of BDNF leads to an impairment of hippocampal-dependent long-term potentiation (LTP), a form of synaptic plasticity, suggesting that BDNF is essential for certain forms of learning and memory (53). Because a brain region-specific signaling profile of CRH was identified, indicating that CRH promotes common signaling pathways in cerebellar and hippocampal but not cortical neurons (36), we suggest that CRH may affect similar processes in the cerebellum. In contrast to CRH, learning and memory impairments are among the most commonly reported behavioral effects of exogenous cannabinoids (for review, see Ref.54). On the other hand, the temporally and spatially restricted release of endocannabinoids facilitates the induction of LTP in the hippocampus in single neurons (55). Because we activated CB1 in all cells by exogenously applied WIN-55,212-2 and CB1 receptors are located on excitatory glutamatergic synapses of granule neurons (56) in our model system, we would rather suggest that there are negative effects on LTP. Moreover, previous work has indicated that induction of cerebellar LTP, which is involved in certain forms of motor learning, including associative eyeblink conditioning and adaptation of the vestibulo-ocular reflex, requires presynaptic Ca2+ influx (57), cAMP production (58), and activation of PKA (59), which are all negatively influenced by CB1 activation. Thus, exogenously applied cannabinoids might mediate their negative effects on learning and memory through dampening the signaling pathway of other neurotransmitter systems that improve cognitive functions and, therefore, regulating the expression of downstream target genes.
Taken together, our study highlights an important role for CRH in regulating BDNF expression. The function of CB1 in inhibiting the CRH signaling cascade is consistent with the role that CB1 ligands generally play. This encompasses the mechanisms by which cannabinoids negatively regulate the cAMP-PKA pathway (46, 49) and also included effects on Ca2+-dependent signaling pathways, which are crucial in the modulation of the expression of the neurotrophins and, in particular, BDNF. The general pharmacological stimulation of CB1 receptors clearly has a negative effect on plasticity (49, 60), which might be mediated by inhibiting signaling pathways, leading to expression changes of target genes. Therefore, further studies should concentrate on the molecular and intracellular interactions of CRH and the endocannabinoid system, not only in terms of stress and glucocorticoid-mediated regulation of the HPA axis, but also in terms of the interactions of these two systems in diverse memory processes with particular attention to the regulation of expression and neuronal release of neurotrophins.
Acknowledgments
We thank Dr. G. Marsicano for statistical evaluation and fruitful discussions and for providing the CB1-deficient mouse line, J. Zschocke and Dr. A. Clement for critical reading of the manuscript, B. W?lfel for excellent support in animal work, and F. Ohl for supplying R121919. CRHR1 plasmid was a kind gift from Dr. W. Wurst.
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