We previously suggested that therapeutic effects of betahistine in vestibular disorders result from its antagonist properties at histamine H3 receptors (H3Rs). However, H3Rs exhibit constitutive activity, and most H3R antagonists act as inverse agonists. Here, we have investigated the effects of betahistine at recombinant H3R isoforms. On inhibition of cAMP formation and [3H]arachidonic acid release, betahistine behaved as a nanomolar inverse agonist and a micromolar agonist. Both effects were suppressed by pertussis toxin, were found at all isoforms tested, and were not detected in mock cells, confirming interactions at H3Rs. The inverse agonist potency of betahistine and its affinity on [125I]iodoproxyfan binding were similar in rat and human. We then investigated the effects of betahistine on histamine neuron activity by measuring tele-methylhistamine (t-MeHA) levels in the brains of mice. Its acute intraperitoneal administration increased t-MeHA levels with an ED50 of 0.4 mg/kg, indicating inverse agonism. At higher doses, t-MeHA levels gradually returned to basal levels, a profile probably resulting from agonism. After acute oral administration, betahistine increased t-MeHA levels with an ED50 of 2 mg/kg, a rightward shift probably caused by almost complete first-pass metabolism. In each case, the maximal effect of betahistine was lower than that of ciproxifan, indicating partial inverse agonism. After an oral 8-day treatment, the only effective dose of betahistine was 30 mg/kg, indicating that a tolerance had developed. These data strongly suggest that therapeutic effects of betahistine result from an enhancement of histamine neuron activity induced by inverse agonism at H3 autoreceptors.
The histamine H3 receptor (H3R) initially was characterized as an autoreceptor regulating histamine release in brain (Arrang et al., 1983). Its coupling to Gi/o proteins was confirmed by its cloning in human (Lovenberg et al., 1999). Activation of recombinant H3Rs inhibits adenylate cyclase (Lovenberg et al., 1999) and activates phospholipase A2 (Morisset et al., 2000a). In different species, including human, various functional isoforms are generated by deletion of a pseudo-intron, variable in length and located in the third intracellular loop of the H3R (Hancock et al., 2003).
Recombinant H3Rs exhibit high constitutive activity, and most antagonists act in fact as inverse agonists on various responses (Arrang et al., 2007). Prototypic antagonists themselves, such as thioperamide or ciproxifan (CPX), act as inverse agonists and abrogate constitutive activity (Morisset et al., 2000a). Moreover, constitutive activity of the recombinant H3R depends on species, isoforms, cell lines, and signaling pathways (Arrang et al., 2007), but all of the data are consistent with high constitutive activity of the rat H3R (rH3R) and human H3R (hH3R).
Consistent with the physiological relevance of the phenomenon, we demonstrated high constitutive activity of native H3Rs in rodent brain in vitro (Morisset et al., 2000a; Rouleau et al., 2002). Moreover, in vivo, H3R-inverse agonists enhance tele-methylhistamine (t-MeHA) levels, a reliable index of histamine neuron activity, by abrogating the brake triggered by constitutive activity of H3 autoreceptors (Morisset et al., 2000a; Gbahou et al., 2003). The recombinant human H3Rs expressed at moderate densities also display constitutive activity, suggesting it is present in human brain (Rouleau et al., 2002).
Betahistine is an orally active drug that has been extensively used in the symptomatic treatment of vestibular disorders such as Meniere's disease (Canty and Valentine, 1981; Oosterveld, 1984). However, the molecular mechanisms underlying its therapeutic effects remain unclear, inasmuch as its first metabolite may also display some activity (Fossati et al., 2001). We have reported previously in the rat that betahistine behaves as a H3R antagonist with moderate potency at autoreceptors modulating histamine release in vitro (Arrang et al., 1985), and we suggested that this property, coupled to moderate H1 receptor agonist activity, might account for the beneficial effects of the compound in the treatment of vertigo. In agreement, the systemic administration of thioperamide results in a strong vestibuloplegic effect and strongly depresses the horizontal vestibular-ocular reflex gain (Yabe et al., 1993). It also accelerates the recovery of behavioral functions in the cat after unilateral vestibular neurectomy (Tighilet et al., 2006, 2007).
The antagonist property of betahistine that we initially reported at H3 autoreceptors (Arrang et al., 1985) was established on histamine release induced by a low stimulus, that is, when the H3 autoreceptor displayed no apparent constitutive activity (Morisset et al., 2000a). However, the improvement of vestibular compensation induced in the cat by betahistine was accompanied by an enhancement of histidine decarboxylase mRNA expression in histamine perikarya (Tighilet et al., 2007), thereby suggesting that betahistine was in fact acting as an H3R inverse agonist.
In the present study, we first investigated the in vitro activity of betahistine at recombinant rH3(413)R and rH3(445)R isoforms mediating inhibition of cAMP formation and [3H]arachidonic acid release in CHO cells. The H3R displays constitutive activity on these two functional responses (Morisset et al., 2000a; Rouleau et al., 2002); therefore, the whole spectrum of drug activity, ranging from agonism to inverse agonism, can be explored. In addition, because H3Rs show important species-related pharmacological differences (Leurs et al., 2005), and betahistine had not yet been studied at human H3Rs, we compared the profiles of betahistine at recombinant human and rat H3(445)Rs in the same tests. We have investigated the in vivo activity of betahistine on histamine neuron activity, by measuring t-MeHA levels, a reliable index of this activity, in the brains of mice after acute intraperitoneal administration and acute or repeated oral administration.
Materials and Methods
Stable Transfection of CHO-K1 Cells.
The clones used in this study were obtained as described previously (Rouleau et al., 2002). cDNA inserts corresponding to the full-length coding sequences of the rH3R and hH3R isoforms were ligated into the mammalian expression vector pCIneo (Promega, Charbonnières, France). CHO-K1 cells were transfected by using SuperFect (QIAGEN S.A., Courtaboeuf, France). Stable transfectants were selected with 2 mg/ml Geneticin (G418) (Invitrogen, Cergy-Pontoise, France), and their expression level was tested for [125I]iodoproxyfan binding, a selective H3R radioligand (Ligneau et al., 1994). Several clones, CHO(rH3(445)R), CHO(rH3(413)R), and CHO(hH3(445)R), expressing receptor densities from 200 to 700 fmol/mg protein were selected for further characterization and maintained in presence of 1 mg/ml G418.
[125I]Iodoproxyfan Binding Assay.
CHO(rH3(445)R), CHO (rH3(413)R), and CHO(hH3(445)R) cells were washed and homogenized with a Polytron homogenizer in ice-cold binding buffer (50 mM Na2HPO4/KH2PO4, pH 7.5). After centrifugation (12,000g for 30 min at 4°C), the pellet was suspended in 1 ml of ice-cold binding buffer, and assays were performed as described previously (Ligneau et al., 1994). Aliquots of membrane suspension (∼20 μg of protein) were incubated for 60 min at 25°C with 20 pM [125I]iodoproxyfan alone (total binding) or together with betahistine at increasing concentrations (200 μl, final volume). The nonspecific binding was determined in the presence of the selective H3R agonist imetit (1 μM). The specific binding was calculated from the difference between the total binding and nonspecific binding and represented ∼80% of the total binding.
CHO(rH3(445)R), CHO(rH3(413)R), and CHO (hH3(445)R) cells expressing 200 to 700 fmol/mg protein were incubated in 96-well plates for 10 min at 37°C with 3 μM forskolin (FSK), the adenylate cyclase activator, and when required, betahistine at increasing concentrations in Dulbecco's modified Eagle's medium-Nut mix F-12 (Invitrogen) containing 100 μM isobutyl-methyl-xanthine. The effects of betahistine were compared with those of histamine (10 μM), the natural agonist, and thioperamide (10 μM), a potent and selective inverse agonist (Morisset et al., 2000a). cAMP was extracted and measured by radioimmunoassay.
[3H]Arachidonic Acid Release.
CHO(rH3(445)R), CHO(rH3(413)R), and CHO(hH3(445)R) cells expressing 200 to 700 fmol/mg protein were incubated for 2 h at 37°C with 0.5 μCi of [3H]arachidonic acid in Dulbecco's modified Eagle's medium-Nut mix F-12 containing 0.2% bovine serum albumin. After washings, cells were incubated for 30 min with 2 μM Ca2+ ionophore A23187 [5-methylamino-2-(2S,3R,5R,8S,9S)-3,5,9-trimethyl-2-(1-oxo-(1H-pyrrol-2-yl)propan-2-yl)-1,7-dioxaspiro-(5,5)undecan-8-yl)methyl)benzooxazole-4-carboxylic acid] and, when required, betahistine at increasing concentrations, histamine, or thioperamide (10 μM). [3H]arachidonic acid release was determined by liquid scintillation counting.
Determination of tele-Methylhistamine Levels in Brain.
Drugs were dissolved in 1% methylcellulose or saline solution [0.9% (w/v) NaCl] for oral and intraperitoneal administration, respectively, to male Swiss mice (18–20g) (Iffa-Credo, L'Arbresle, France). After treatment, animals were sacrificed by decapitation. The total brain was dissected out and homogenized in 10 volumes (w/v) of ice-cold perchloric acid (0.4 N). The clear supernatant was stored at −20°C immediately after centrifugation (4,000g for 20 min). t-MeHA levels were determined by using an enzyme immunoassay derived from a radioimmunoassay described previously (Garbarg et al., 1989a, 1992). In brief, t-MeHA of the sample was derivatized with p-benzoquinone (BZQ) (2.8 mg/ml). The reaction was allowed to proceed at pH 7.9 for 3 h, then 2 M glycine was added to eliminate the excess of BZQ. The derivatized extract was mixed with t-MeHA-BZQ-Leu-Tyr-acetylcholinesterase as a tracer and an antiserum raised in rabbits against t-MeHA conjugated with bovine serum albumin via p-benzoquinone in a plate (Nunc Immuno-Plate Maxi-Sorp Surface; NUNC A/S, Roskilde, Denmark) pretreated with swine anti-rabbit IgG (Cayman Chemical, Ann Arbor, MI). After incubation for 16 h at 15°C, plates were washed, and the substrate for acetylcholinesterase, the Ellmann's reagent, was added. After 5 h, the optical density was measured with a Dynatech MR5000 (Dynex Technologies, Chantilly, VA) at 405 nm. The limit of the detection was 5 pg of t-MeHA.
Analysis of Data.
In binding experiments, IC50 values of betahistine were determined with an iterative least-squares method derived from that of Parker and Waud (1971). The apparent affinity constants (Ki values) of betahistine were calculated from its IC50 values, assuming a competitive antagonism and by using the relationship (Cheng and Prussoff, 1973) Ki = IC50/1 + (S/KD), where S represents the concentration (20 pM) and KD is the apparent dissociation constant of [125I]iodoproxyfan at the rH3(445)R (85 ± 4 pM) and hH3(445)R (82 ± 3 pM).
In functional assays, EC50 values of betahistine corresponding to stimulatory and inhibitory phases of its biphasic effects were evaluated by nonlinear regression with Prism software (GraphPad Software, Inc., San Diego, CA). The following equation was fitted to the bell-shaped concentration-response curves: Y = Dip + Section1 + Section2, where Y = the observed response in the presence of a given concentration of betahistine, expressed as percentage of the response in its absence. Dip is plateau level between phases. Section1 = Span1/(1 + 10((logEC50_1 − X) × nH1)), and Section2 = Span2/(1 + 10((X −logEC50_2) × nH2)), where Span1 = Plateau1 − Dip and Span2 = Plateau2 − Dip, Plateau1 and Plateau2 are initial and final plateau levels, respectively, X is log of the concentration of betahistine (M), logEC50_1 and logEC50_2 are logEC50 values for respective phases, and nH1 and nH2 are slope factors for respective phases.
Protein content was determined according to the method of Lowry et al. (1951), using bovine serum albumin as the standard. Statistical evaluation of the results was performed by one-way analysis of variance followed by Newman-Keuls test.
Radiochemicals and Drugs.
[125I]iodoproxyfan (2000 Ci/mmol) was prepared as described previously (Krause et al., 1997). [3H]arachidonique acid (211 Ci/mmol) was from PerkinElmer Life and Analytical Sciences (Waltham, MA). Isobutyl-methyl-xanthine, pertussis toxin (PTX), forskolin, and bovine serum albumin were purchased from Sigma-Aldrich (Saint Quentin Fallavier, France). Ionophore A23187 was obtained from Roche Diagnostics (Meylan, France). (R)-α-methylhistamine [(R)-α-MeHA], thioperamide, and ciproxifan were from Bioprojet (Paris, France). Betahistine was from Solvay Pharma (Suresnes, France).
Effect of Betahistine on [125I]Iodoproxyfan Binding to Rat and Human H3R Isoforms.
Computer analysis by nonlinear regression using a one-site cooperative model showed that betahistine inhibited [125I]iodoproxyfan binding to membranes of CHO(rH3(445)R) and CHO(hH3(445)R) cells with pseudo-Hill coefficients not significantly different from unity. IC50 values of 1.9 ± 0.2 and 3.3 ± 0.4 μM, respectively, led to Ki values of betahistine of 1.4 ± 0.1 μM at the rH3(445)R and 2.5 ± 0.3 μM at the hH3(445)R (Fig. 1; Table 1). Betahistine inhibited the binding of [125I]iodoproxyfan to membranes of CHO(rH3(413)R) cells with a pseudo-Hill coefficient of 0.5 ± 0.1 and a mean IC50 value of 9 ± 2 μM (Fig. 1; Table 1). The data fitted significantly better to a two-site model analysis and could be resolved in a high-affinity site with an IC50 value of 0.9 ± 0.3 nM and a low-affinity site with an IC50 value of 77 ± 42 μM.
Functional Properties of Betahistine at Rat H3R Isoforms.
In agreement with the H3R-mediated inhibition of adenylyl cyclase, histamine used as an agonist significantly inhibited cAMP formation induced by 3 μM FSK in CHO(rH3(445)R) cells, with an EC50 value of 9 ± 2 nM (not shown) and a maximal effect reached at 1 to 10 μM of −20 ± 2% (n = 9 experiments). In contrast, the reference inverse agonist thioperamide (10 μM) enhanced FSK-induced-cAMP accumulation in CHO(rH3(445)R) cells (+35 ± 8%; n = 7 experiments), revealing the constitutive activity of the rat H3(445)R isoform (Fig. 2).
Increasing concentrations of betahistine promoted a biphasic dose–response curve in CHO(rH3(445)R) cells. When added at low concentrations, betahistine mimicked the effect of thioperamide, thereby behaving as an apparent inverse agonist, and progressively enhanced cAMP formation (EC50 = 0.1 ± 0.1 nM) with a maximal effect, observed up to 10 nM, similar to that of thioperamide (+32 ± 7% versus + 35 ± 8%; p < 0.001 compared with FSK alone). In contrast, at concentrations higher than 10 nM betahistine progressively inhibited cAMP formation with an EC50 value of 0.1 ± 0.1 μM and full agonist activity, its maximal effect being similar to that of histamine (−21 ± 3% versus −20 ± 2%; p < 0.001 versus FSK alone) (Fig. 2, A and B; Table 1). This agonist effect was entirely blocked by 10 μM thioperamide (4.5 ± 0.2 pmol cAMP versus 9.5 ± 0.8 pmol cAMP, respectively; p < 0.001).
In agreement with the H3R-mediated activation of phospholipase A2, histamine used as an agonist at a maximal concentration (10 μM) strongly enhanced [3H]arachidonic acid release evoked by the Ca2+ ionophore A23187 from CHO(rH3(445)R) cells (+48 ± 6%; n = 19 experiments). As already observed in rat (Morisset et al., 2000a; Gbahou et al., 2003), its EC50 value on this response was higher than on cAMP formation (141 ± 39 nM versus 9 ± 2 nM; not shown). In contrast, the inverse agonist thioperamide (10 μM) decreased significantly A23187-evoked [3H]arachidonic acid release (−44 ± 4%; n = 22 experiments), revealing constitutive activity of the rat H3(445)R isoform in this test system. A biphasic response was observed at the rat H3(445)R in the presence of increasing concentrations of betahistine. At low concentrations, betahistine progressively reduced A23187-evoked [3H]arachidonic acid release (EC50 = 0.1 ± 0.1 nM) with a maximal effect, observed up to 30 nM, similar to that of thioperamide (−47 ± 5 and −44 ± 4%, respectively; p < 0.001 versus A23187 alone). At concentrations higher than 30 nM, betahistine progressively enhanced the release with an EC50 value of 7 ± 1 μM and full agonist activity, its maximal effect being similar to that of histamine (+41 ± 11% versus +48 ± 6%; p < 0.01 versus A23187 alone) (Fig. 2, C and D; Table 1). At this rat isoform, betahistine therefore had the same inverse agonist potency on cAMP formation and [3H]arachidonic acid release (EC50 of 0.1 ± 0.1 and 0.1 ± 0.1 nM, respectively), whereas its agonist potency, like that of histamine, was lower on [3H]arachidonic acid release (EC50 of 7 ± 1 μM versus 0.1 ± 0.1 μM; Table 1).
In CHO cells expressing the short rat H3(413)R isoform, the same biphasic patterns were observed, both on cAMP accumulation and [3H]arachidonic acid release (Fig. 3). Betahistine increased in a dose-dependent manner the concentration of cAMP accumulation (EC50 = 0.05 ± 0.15 nM) to reach a maximal effect, observed up to 1 nM, similar to that of thioperamide (+17 ± 4% versus + 22 ± 2%,;p < 0.05 versus FSK alone) (Fig. 3, A and B; Table 1). At concentrations higher than 1 nM betahistine behaved as histamine and inhibited cAMP accumulation by 73% at the highest concentration tested (300 μM). An EC50 value ≥15 ± 1 μM was obtained for betahistine acting as an agonist at CHO(rH3(413)R) cells, assuming that the drug was acting as a full agonist (Table 1). This agonist effect was again entirely blocked by 10 μM thioperamide (3.9 ± 0.7 pmol cAMP versus 11.9 ± 1.2 pmol cAMP, respectively; p < 0.001).
On [3H]arachidonic acid release from CHO(rH3(413)R) cells betahistine displayed a similar biphasic effect. It behaved as a full inverse agonist up to 3 nM (−16 ± 5% versus −20 ± 2% for thioperamide; p < 0.05 versus A23187 alone). Its agonist effect reached a plateau corresponding to 51% of the maximal effect of histamine at 100 μM, suggesting that betahistine was acting as a partial agonist (Fig. 3, C and D). EC50 values of 0.06 ± 0.23 nM and 4 ± 1 μM were obtained on this response at CHO(rH3(413)R) cells for betahistine acting as an inverse agonist and partial agonist, respectively (Table 1). At this short rat isoform, betahistine had a similar potency on cAMP formation and [3H]arachidonic acid release, not only as an inverse agonist (EC50 of 0.05 ± 0.15 and 0.06 ± 0.23 nM, respectively), but also as an agonist (EC50 ≥ 15 ± 1 and 4 ± 1 μM, respectively), in contrast to histamine itself, which remained less potent on [3H]arachidonic acid release than on cAMP formation at this isoform (EC50 of 110 ± 32 nM versus 15 ± 4 nM; not shown).
Functional Properties of Betahistine at the Human H3(445)R Isoform.
Histamine at a maximal concentration (1–10 μM) significantly inhibited cAMP formation induced by 3 μM FSK in CHO(hH3(445)R) cells (−91 ± 1%; n = 9 experiments). In contrast, thioperamide (10 μM) enhanced FSK-induced-cAMP accumulation in CHO(hH3(445)R) cells (+20 ± 4%; n = 9 experiments), revealing the constitutive activity of the human H3(445)R (Fig. 4). Increasing concentrations of betahistine promoted the biphasic dose–response curve shown at the recombinant rat H3R isoforms (Fig. 4, A and B). When added at low concentrations, betahistine mimicked the effect of thioperamide, thereby behaving as an apparent inverse agonist, and progressively enhanced cAMP formation (EC50 = 0.3 ± 1.4 nM) with a maximal effect, observed up to 0.1 μM, similar to that of thioperamide (+15 ± 4% versus +20 ± 4%; p < 0.05 versus FSK alone). In contrast, at concentrations higher than 0.1 μM the effect of betahistine was similar to that observed for histamine, with a progressive inhibition of cAMP formation that reached 70% at the highest concentration tested (300 μM) (Fig. 4B). An EC50 value ≥48 ± 2 μM was obtained, assuming that betahistine was acting as a full agonist (Table 1). As expected, this agonist effect was entirely blocked by 10 μM thioperamide, because thioperamide tested against betahistine increased cAMP formation with an amplitude (+22%) similar to that observed when it was used alone (Fig. 5A).
The effects of betahistine on cAMP formation were not observed in CHO cells transfected with the empty vector (mock cells) (Fig. 5B). In another set of experiments, the effects of betahistine on cAMP formation were studied at its highest doses as inverse agonist (1 nM) and agonist (100 μM) without or with PTX. In the absence of PTX the biphasic effect of betahistine was observed (Fig. 5C), but both the inverse agonist and the agonist effects of betahistine were completely abolished in the presence of PTX (Fig. 5D).
On [3H]arachidonic acid release, histamine used as an agonist at a maximal concentration (1–10 μM) strongly enhanced [3H]arachidonic acid release evoked by the Ca2+ ionophore A23187 from CHO(hH3(445)R) cells (+278 ± 39%; n = 19 experiments). In contrast, the inverse agonist thioperamide (10 μM) significantly decreased A23187-evoked [3H]arachidonic acid release (−31 ± 3%; n = 21 experiments), revealing constitutive activity of the human H3R in this test system. Increasing concentrations of betahistine promoted a biphasic dose–response curve in CHO(hH3(445)R) cells (Fig. 4, C and D; Table 1). When added at low concentrations, betahistine behaved as an apparent inverse agonist and progressively decreased [3H]arachidonic acid release (EC50 = 0.1 ± 0.3 nM) with a maximal effect, observed up to 30 nM, similar to that of thioperamide (−25 ± 3% versus −31 ± 3%; p < 0.01 versus A23187 alone). In contrast, at concentrations higher than 30 nM, the effect of betahistine on CHO(hH3(445)R) cells was similar to that observed for histamine, with a progressive increase in A23187-evoked [3H]arachidonic acid release that reached 180 ± 31% at the highest concentration tested (300 μM). An EC50 value ≥65 ± 20 μM was obtained for betahistine acting as an agonist, assuming that the drug was acting as a full agonist. The stimulation induced by betahistine was entirely blocked by a maximal dose of thioperamide, and thioperamide tested against betahistine decreased [3H]arachidonic acid release to the same amplitude as that observed when it was used alone (4452 ± 485 dpm versus 3172 ± 327 dpm; p < 0.05).
Effects of Intraperitoneal or Oral Administration of Betahistine on tele-Methylhistamine Levels in Mouse Brain.
A single intraperitoneal administration of the inverse agonist ciproxifan, used at the maximal dose of 3 mg/kg as a control, enhanced t-MeHA levels by 89 ± 10% (238 ± 10 versus 126 ± 8 ng/g; n = 20–22 mice). The acute intraperitoneal administration of betahistine increased t-MeHA levels in a dose-dependent manner (Fig. 6). At low doses, t-MeHA levels progressively increased (ED50 = 0.4 ± 0.1 mg/kg) to reach a peak of ∼+30% (p < 0.05) after administration of 1 mg/kg. After intraperitoneal administration of higher doses of betahistine, the enhancement became gradually lower with t-MeHA levels returning to control values at 10 to 30 mg/kg (Fig. 6).
A single oral administration of ciproxifan (3 mg/kg) enhanced t-MeHA levels by 117 ± 28% (215 ± 22 versus 99 ± 17 ng/g; n = 8 mice). The acute oral administration of betahistine increased t-MeHA levels in a dose-dependent manner (Fig. 7) with an ED50 of 2 ± 1 mg/kg and a maximal effect of ∼35% reached at 30 mg/kg.
Repeated oral administration of ciproxifan (3 mg/kg/day for 8 days) was followed 90 min after the last administration by a significant increase (+55%) of brain t-MeHA levels (Fig. 7). Repeated oral administration of the standard agonist (R)-α-MeHA (3 mg/kg/day for 8 days) did not change t-MeHA levels. Repeated oral administration of betahistine at 3 mg/kg/day for 8 days had no significant effect, but betahistine at 30 mg/kg/day for 8 days significantly increased t-MeHA levels by 37 ± 13% (Fig. 7).
At recombinant H3Rs betahistine behaved not only as a potent inverse agonist, but also as an agonist at higher concentrations. These inverse agonist and agonist effects both were mediated by H3Rs. They were found on a negatively (cAMP accumulation) and positively ([3H]arachidonic acid release) coupled response. They were not detected in mock cells, but both were found in all rat and human isoforms. They were both suppressed by PTX, in agreement with coupling of H3Rs to Gi/o proteins. This wide spectrum of activity did not result from protean agonism (Kenakin, 1995), because, in contrast to proxyfan (Gbahou et al., 2003), it occurred at the same level of constitutive activity. It did not result from a switch in coupling from Gi to Gs proteins, such as that reported with agonists at muscarinic receptors (Michal et al., 2001), because opposite effects of betahistine would have been observed. Moreover, their PTX sensitivity shows that the two responses are mediated by Gi/Go proteins. Further studies are required to know which Gi/Go proteins are involved, inasmuch as the hH3(445)R couples similarly to different Gαi/o subunits (Schnell et al., 2010).
The multistate model of G protein-coupled receptors (Kenakin, 1995; Gbahou et al., 2003; Arrang et al., 2007) more likely accounts for these biphasic effects. Betahistine would first stabilize inactive conformations, thereby inducing inverse agonism. At higher concentrations, betahistine would stabilize an active conformation antagonized by thioperamide. To our knowledge, betahistine is so far the first H3R ligand inducing such biphasic effects, which suggests some ligand selectivity. Conformations selective for a given agonist, and with preferential coupling properties, have been suggested for various G protein-coupled receptors (Kenakin, 2001). Although their existence remains controversial for the H3R (Krueger et al., 2005; Schnell et al., 2010), a selective betahistine-directed state would account for the trafficking from inverse agonism to agonism. It may also account for the large variations of agonist potencies of betahistine at rat isoforms: the preferential negative coupling of the long isoform to adenylate cyclase induced by histamine and betahistine, was maintained for histamine, but not for betahistine, at the short isoform.
The inverse agonist potency of betahistine was surprisingly high, compared with its affinity on [125I]iodoproxyfan binding. However, inverse agonist radioligands label a much larger population of H3Rs than agonist radioligands such as [125I]iodoproxyfan (Witte et al., 2006; Yao et al., 2006; Mezzomo et al., 2007). Therefore, inactive conformations with high affinity for betahistine are not expected to be labeled by [125I]iodoproxyfan. Nevertheless, betahistine displaced [125I]iodoproxyfan binding to the short rH3(413)R isoform with a Hill coefficient lower than unity. Although we did not analyze the effect of GTP on this shallow inhibition curve, it may reflect binding of betahistine to both uncoupled and coupled agonist states of this short isoform (Arrang et al., 1990). Moreover, the mean IC50 of betahistine at this rH3(413)R isoform was higher than its Ki at the rH3(445)R isoform (Table 1), but very similar to its Ki at rat autoreceptors (Arrang et al., 1985), suggesting that the function of autoreceptor is fulfilled by short, rather than long, H3R isoforms.
In contrast to other antagonists/inverse agonists (Leurs et al., 2005), the inverse agonist potency and binding affinity of betahistine were similar in rat and human. However, its agonist potencies on inhibition of cAMP formation and [3H]arachidonic acid release were lower at the human H3R (Table 1). So far, betahistine is the first agonist to discriminate the human and rat receptors (Wulff et al., 2002). In addition, the preferential negative coupling to adenylate cyclase induced in rat by betahistine was no longer observed in human. Its binding affinity remaining unchanged at the two H3Rs. The betahistine selective state discussed above may promote a differential coupling not only of different isoforms from a given species, but also of a given isoform from different species.
Consistent with its inverse agonist potency, a single intraperitoneal administration of betahistine increased t-MeHA levels. However, its maximal effect was lower than that of ciproxifan, indicating partial inverse agonism. At higher doses, t-MeHA levels returned to basal levels, yielding a bell-shaped curve reminiscent of biphasic curves in vitro. This suggests that H3R agonism progressively reversed inverse agonism in vivo. It is noteworthy that betahistine also acted as an agonist to increase cochlear blood flow in the guinea pig (Laurikainen et al., 1998). The contribution of H1 receptors activated by betahistine to this bell-shaped curve can be excluded, because they are postsynaptic and do not regulate histamine neuron activity (Garbarg et al., 1989b).
In contrast to ciproxifan, the profile of betahistine on histamine neuron activity became different when it was given orally. It acted only as a partial inverse agonist and at doses ∼10 times higher. This rightward shift probably is caused by the rapid metabolism of betahistine in the liver after oral administration. Very low plasma concentrations were found after its oral administration to human volunteers, indicating that it undergoes almost complete first-pass metabolism (Chen et al., 2003). The rightward shift of its ED50 shows that only 10% of the initial dose remains in tissues 90 min later, explaining that agonism does not occur in such conditions, even at the highest doses.
As expected (Morisset et al., 2000b), repeated administration of ciproxifan induced a tolerance, its effect at 3 mg/kg being reduced by half after 8-day oral administration. After 8 days, the increase induced by oral administration of betahistine was only found at the high dose of 30 mg/kg, suggesting either that a tolerance had developed for betahistine as an inverse agonist or that betahistine at such a high dose was acting as an agonist. Theoretically, repeated administration of an agonist may desensitize autoreceptors, relieve the brake, and lead to an enhancement of t-MeHA levels. (R)-α-methylhistamine itself induced a desensitization, because its repeated administration failed to decrease histamine neuron activity. However, it also failed to enhance t-MeHA levels, confirming that the increase induced by repeated administration of betahistine at 30 mg/kg was not caused by desensitization of autoreceptors by its agonist effect. Therefore, metabolism and tolerance made a high dose necessary to induce inverse agonism after repeated oral administration of betahistine.
Overall, our data show that betahistine acts as an inverse agonist to increase histamine neuron activity. In agreement, Tighilet et al. (2002, 2005) reported that betahistine given orally to cats increased histidine decarboxylase mRNA expression in the tuberomammilllary nucleus. Moreover, in mice (the present study) and cats, the increase in histamine neuron activity was observed after a 1- to 3-week treatment, i.e., when lesioned animals treated by betahistine recover their vestibular functions (Tighilet et al., 2007).
The present study confirms that human H3Rs display constitutive activity (Rouleau et al., 2002). However, whether inverse agonism and enhancement of histamine neuron activity account, even partially, for the therapeutic effects of betahistine requires further investigations, including positron emission tomography studies. After 1-week oral administration, the effective doses of betahistine in mice were higher than the therapeutic dose (2 mg/kg) given, generally twice a day, to vestibular-defective patients. However, the dose–response curve of betahistine on histamine neuron activity in lesioned animals is unknown. Its ED50 may be lower than in controls, because imbalanced vestibular activity itself activates histamine neurons (Tighilet et al., 2006, 2007) and the effect of the drug may synergize with this compensatory activation. Moreover, in clinical practice, this therapeutic dose of 2 mg/kg is given for several months. In agreement with our findings, Tighilet et al. (2006, 2007) reported that this dose had no effect on histidine decarboxylase mRNA expression in control cats after a 1- or 3-week treatment. However, the same low dose acquired an enhancing effect after 3 months (Tighilet et al., 2005). This suggests either that betahistine progressively accumulates into the brain or simply that a low dose given over a long period of time yields the therapeutic effect. Its activation of histamine neurons indicates that betahistine enters the rodent brain after systemic administration, even though its effective doses in vivo are rather high compared with its subnanomolar inverse agonist potency in vitro. In addition, the effect of betahistine may be enhanced and prolonged in time by its first metabolite [(2–2-aminoethyl)-pyridine], which has the same affinity at rodent H3R binding sites (Fossati et al., 2001). Although we did not investigate the functional properties of this metabolite, it should be emphasized that its presence, presumably predominant after oral and repeated administration, renders even more complex the interpretation of in vivo and therapeutic effects of betahistine.
In conclusion, betahistine interacts in vitro with H3Rs as a potent inverse agonist, a moderate antagonist (Arrang et al., 1985), and a weak agonist. It acts in vivo as a partial inverse agonist to enhance histamine neuron activity, an effect probably involved in therapeutics. Therefore, although H3 heteroreceptors present on other neurons regulating vestibular functions may also be involved (Lozada et al., 2004; Chavez et al., 2005; Bergquist et al., 2006), this study strongly suggests that H3 autoreceptors regulating histamine neurons in human brain constitute a major target under betahistine treatment.
We thank M. O. Christen and B. Laroche for scientific collaboration and Dominique Dumoulin for excellent technical assistance.
This study was supported by Institut National de la Santé et de la Recherche Médicale, the Fondation pour la Recherche Médicale, and the Direction des Systèmes de Forces et de la Prospective.
Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
- H3 receptor
- human H3R
- rat H3R
- pertussis toxin
- Chinese hamster ovary
- 5-methylamino-2-(2S,3R,5R,8S,9S)-3,5,9-trimethyl-2-(1-oxo-(1H-pyrrol-2-yl)propan-2-yl)-1,7-dioxaspiro-(5,5)undecan-8-yl)methyl)benzooxazole-4-carboxylic acid.
- Received March 25, 2010.
- Accepted June 2, 2010.
- Copyright © 2010 by The American Society for Pharmacology and Experimental Therapeutics