Animals

All animal care and experimental procedures complied with Italian (D.L. 116/92) and EEC (O.J. of EC L358/1 18/12/1986) regulations on the protection of laboratory animals, and with the UK Animals (Scientific Procedures) Act 1986 and associated guidelines for the use of experimental animals. Guidelines of the International Association for the Study of Pain were also followed. Brain and spleen tissue was obtained from adult male C57BL/6J mice weighing 25–40 g and maintained on a 12/12 h light/dark cycle with free access to food and water. These animals were either wild-type mice (Harlan UK Ltd, Blackthorn, UK) or mice from which the CB₂ receptor had been genetically deleted as described by Buckley et al., 2000. All in vivoexperiments were also performed with male C57BL/6J mice (Harlan, Milan, Italy). These mice were 9 weeks old and housed three per cage under controlled illumination (12:12 h light : dark cycle; light on at 0600 h) and standard environmental conditions (room temperature 22 ± 1°C; humidity 60 ± 10%) for at least 1 week before experimental use. Mouse chow and tap water were available ad libitum. All the in vivo experiments were performed in a randomized manner by an experimenter, unaware of the pharmacological treatments.

CHO cells

CHO cells either untransfected or transfected with cDNA encoding human cannabinoid CB₂ or CB₁ receptors were maintained in Dulbecco’s modified Eagle’s medium nutrient mixture F-12 HAM, supplemented with 1 mM l-glutamine, 10% fetal bovine serum and 0.6% penicillin–streptomycin for all cells together with G418 (400 µg·mL⁻¹) for the CHO–hCB₂ cells or with hygromycin B (300 µg·mL⁻¹) and G418 (600 µg·mL⁻¹) for the CHO–hCB₁cells. All cells were maintained at 37°C and 5% CO₂ in their respective media, and were passaged twice a week using non-enzymatic cell dissociation solution.

Membrane preparation

Binding assays with [³H]CP55940 or [³⁵S]GTPγS were performed with CHO–hCB₂ cell membranes (Ross et al., 1999a), with mouse whole brain membranes (Viganòet al., 2003) or with mouse spleen membranes, the preparation of which was based on a method described by Hillard et al. (1999) for preparing rat spleen membranes. The hCB₂ receptor-transfected cells were removed from flasks by scraping and then frozen as a pellet at −20°C until required. Before use in a radioligand binding assay, cells were defrosted, diluted in Tris buffer (50 mM Tris–HCl and 50 mM Tris–base) and homogenized with a 1 mL hand-held homogenizer. Protein assays were performed using a Bio-Rad DC Kit (Hercules, CA, USA). Spleens were cut into several pieces and placed in a Choi lysis buffer (Tris–HCl 20 mM, sucrose 0.32 M, EDTA 0.2 mM, EGTA 0.5 mM, pH 7.5) containing Roche protease inhibitor cocktail (1:40 v/v; Roche Diagnostics, Mannheim, Germany) and phenylmethylsulphonyl fluoride (PMSF; 150 µM), and then homogenized. The homogenate was centrifuged at 500×g for 2 min, and the resulting supernatant was re-centrifuged at 16 000×g for 20 min. The harvested membranes were resuspended in TME buffer (50 mM Tris–HCl; EDTA 1.0 mM; MgCl₂ 3.0 mM; pH 7.4) and stored at −80°C for no more than 1 month. Mouse brains were homogenized in ice-cold Choi lysis buffer containing Roche protease inhibitor cocktail (1:40 v/v) and PMSF (1 mM). The homogenate was centrifuged at 13 500×g for 15 min, and the resulting pellet was kept at −80°C for at least 2 h. The pellet was then resuspended in TME buffer, homogenized and stored at −80°C.

Cyclic AMP assay

Adherent CHO–hCB₁ or CHO–hCB₂ cells were washed once with Dulbecco’s phosphate-buffered saline (PBS) and detached using non-enzymatic cell dissociation solution. After centrifugation, cells were resuspended (2 × 10⁶cells·mL⁻¹) in buffer containing PBS (calcium and magnesium free), 1% BSA and 10 µM rolipram. Cells were incubated for 30 min at 37°C with the cannabinoid under investigation. A further 30 min incubation was carried out with 10 µM of forskolin in a total volume of 500 µL. The reaction was terminated by the addition of 0.1 M HCl, followed by centrifugation to remove cell debris. The pH was then adjusted to between 8 and 9 by the addition of 1 M of NaOH, and cyclic-AMP content was measured using a radioimmunoassay kit (GE Healthcare Amersham Ltd, Little Chalfont, Buckinghamshire, UK). Forskolin and rolipram were dissolved in dimethyl sulphoxide (DMSO) and stored at −20°C as 10 mM stock solutions. Some CHO cells were pretreated overnight with pertussis toxin (100 ng·mL⁻¹; Couttset al., 2001). The pertussis toxin used in these experiments was dissolved in distilled water and stored at 4°C.

Radioligand displacement assay

The assays were carried out with [³H]CP55940 and Tris-binding buffer (50 mM Tris–HCl, 50 mM Tris–base, 0.1% BSA, pH 7.4), total assay volume 500 µL, using the filtration procedure described previously by Ross et al. (1999b). Binding was initiated by the addition of transfected hCB₂ cells (50 µg protein per well). All assays were performed at 37°C for 60 min before termination by the addition of ice-cold Tris-binding buffer and vacuum filtration using a 24-well sampling manifold (Brandel Cell Harvester; Brandel Inc, Gaithersburg, MD, USA) and Brandel GF/B filters that had been soaked in wash buffer at 4°C for at least 24 h. Each reaction well was washed six times with a 1.2 mL aliquot of Tris-binding buffer. The filters were oven-dried for 60 min and then placed in 5 mL of scintillation fluid (Ultima Gold XR, PerkinElmer, Seer Green, Buckinghamshire, UK). Radioactivity was quantified by liquid scintillation spectrometry. Specific binding was defined as the difference between the binding that occurred in the presence and absence of 1 µM unlabelled CP55940. The concentration of [³H]CP55940 used in our displacement assays was 0.7 nM. The compounds under investigation were stored as stock solutions of 10 mM in DMSO, the vehicle concentration in all assay wells being 0.1% DMSO. The binding parameters for [³H]CP55940 were 215 pmol·mg⁻¹ (B_max) and 4.3 nM (K_d).

[³⁵S]GTPγS binding assay

The method used for measuring agonist-stimulated binding of [³⁵S]GTPγS was based on previously described methods (Hillard et al., 1999; Thomas et al., 2005). The assays were carried out with GTPγS binding buffer (50 mM Tris–HCl, 100 mM NaCl, 0.1% BSA) in the presence of [³⁵S]GTPγS and GDP, in a final volume of 500 µL. The GTPγS binding buffer also contained 50 mM Tris–base, 5 mM MgCl₂, 1 mM dithiothreitol and 1 mM EDTA (CHO cell membrane experiments) or 3 mM MgCl₂ and 0.2 mM EGTA (brain and spleen membrane experiments). Binding was initiated by the addition of [³⁵S]GTPγS to the wells. Non-specific binding was measured in the presence of 30 µM GTPγS. The drugs were incubated in the assay for 60 min at 30°C. The reaction was terminated by a rapid vacuum filtration method using Tris-binding buffer, and the radioactivity was quantified by liquid scintillation spectrometry. In all the [³⁵S]GTPγS-binding assays, we used 0.1 nM [³⁵S]GTPγS, 30 µM GDP and 10 µg (brain membranes), 40 µg (spleen membranes) or 50 µg (cell membranes) protein per well. Additionally, mouse brain and spleen membranes were pre-incubated for 30 min at 30°C with 0.5 U·mL⁻¹ adenosine deaminase (200 U·mL⁻¹) to remove any endogenous adenosine.

Carrageenan-induced inflammation

The mice were anaesthetized with sodium pentobarbital (60 mg·kg⁻¹ i.p.). Acute inflammation was induced by intraplantar injection of 20 µL of λ-carrageenan (2% w/v in saline) into the right hind paw. The volume of the injected paw, as well as of the contralateral paw, was measured with a plethysmometer (Ugo Basile, Varese, Italy). Data are expressed as oedema (difference in volume between the right and left paws). Responses to thermal stimuli were measured in the same animals used to monitor oedema. After recording baseline withdrawal latencies (s), withdrawal latencies of both hind paws were estimated at different time-points after carrageenan injection, starting 3 h after injection of this inflammatory stimulus, when all mice had recovered from anaesthesia. Heat hypersensitivity was tested according to the Hargreaves procedure (Hargreaves et al., 1988) using the plantar test (Ugo Basile). Briefly, the animals were placed in a clear plexiglass box and allowed to acclimatize. A constant intensity radiant heat source was aimed at the midplantar area of the hind paw. The time, in seconds, from initial heat source activation until paw withdrawal was recorded. The control animals received saline instead of carrageenan by intraplantar injection. All these experiments were performed by the same experimenter.

Formalin test

Formalin injection induces biphasic stereotypical nocifensive behaviour (Dubuisson and Dennis, 1977). Nociceptive responses are divided into an early, short lasting first phase (0–7 min) caused by a primary afferent discharge produced by the stimulus and a subsequent second, prolonged phase (15–60 min) of tonic pain (Sawynok and Liu, 2004). These two phases are separated by a transient quiescent period. The mice received formalin (1.25% in saline, 30 µL) into the dorsal surface of one side of the hind paw. Each mouse was randomly assigned to one of the experimental groups and placed in a plexiglass cage, and allowed to move freely for 15–20 min. A mirror was placed at a 45° angle under the cage to allow full view of the hind paws. Lifting, favouring, licking, shaking and flinching of the injected paw were recorded as nociceptive responses (Abbott and Guy, 1995). Nociceptive responses were measured every 5 min and expressed as their total duration in min (mean ± SEM). Recording of nociceptive behaviour commenced immediately after formalin injection and was continued for 60 min. All these experiments were performed by the same experimenter.

In vivo drug treatments and experimental design

Different doses of THCV or an appropriate volume of its vehicle were administered i.p., 30 min before carrageenan injection or 15 min before formalin injection. THCV was dissolved in a mixture of ethanol : cremophorEL : saline (1:1:18), when tested in the carrageenan model, or in 0.5% DMSO in saline (0.9% aqueous solution of NaCl), when tested using the formalin test. These vehicles were chosen because they have previously been shown to be devoid of activity in the carrageenan and formalin models, respectively (Costaet al., 2004; Maione et al., 2007). Carrageenan-induced oedema and thermal hyperalgesia were evaluated 2 and 3 h after the injection of phlogogen respectively. The maximal effective dose (0.3 mg·kg⁻¹ i.p.) was employed to evaluate the time-course of the anti-inflammatory effects of THCV: oedema was measured at 2, 3, 4, 6 and 24 h and thermal hyperalgesia at 3, 6 and 24 h after carrageenan administration. Some experiments were performed with the selective CB₁ receptor antagonist/inverse agonist, rimonabant (0.5 mg·kg⁻¹), or with the selective CB₂ receptor antagonist/inverse agonist, SR144528 (1 mg·kg⁻¹). These were administered to the mice i.p. 15 min before THCV in both carrageenan and formalin experiments. Oedema and withdrawal latency were evaluated at 2 and 3 h after carrageenan injection, and nociceptive behaviour was observed for 60 min after formalin administration. Rimonabant and SR144528 were dissolved in a mixture of Tween80 : DMSO : distilled water (1:2:7). To determine the ability of THCV to improve established signs of inflammation and pain, an additional series of experiments was performed in which this compound was administered to the mice at 0.3 mg·kg⁻¹ i.p. 30 min after intraplantar injection of carrageenan, and for 3 consecutive days after the induction of inflammation. Behavioural evaluations were performed 2, 24, 48 and 72 h after carrageenan injection. No overt behavioural changes were observed in this study following administration of vehicle or of any of the drugs at the dosage used. The mice remained alert and generally active throughout these experiments.

Statistical analysis

Values have been expressed as means and variability as SEM or as 95% confidence intervals (CIs). The concentration of THCV and CP55940 that produced a 50% displacement of [³H]CP55940 from specific binding sites (IC₅₀ value) was entered into the equation of Cheng and Prusoff (1973) to calculate the corresponding K_i value. Values obtained in vitro for EC₅₀ and maximal effect (E_max) have been calculated by non-linear regression analysis using the equation for a sigmoid concentration–response curve. Statistical analysis of all in vivo data was performed by anova followed by the Newman–Keuls multiple comparison test. All these statistical analyses were performed using GraphPAD software (San Diego, CA, USA). Differences were considered significant at P < 0.05.

Materials

THCV, extracted from Cannabis, was a gift from GW Pharmaceuticals (Porton Down, Wiltshire, UK). It was 99.4% pure and contained no detectable Δ⁹-tetrahydrocannabinol, cannabinol or cannabidiol. Rimonabant and SR144528 were kindly supplied by Sanofi-Aventis (Montpellier, France). Carrageenan and formalin were purchased from Sigma-Aldrich (Milan, Italy) and (–)-cis-3-[2-hydroxy-4-(1,1-dimethylheptyl)phenyl]-trans-4-(3-hydroxypropyl)cyclohexanol (CP55940), from Tocris (Bristol, UK). Pertussis toxin, forskolin and rolipram were supplied by Sigma-Aldrich (Poole, Dorset, UK) and [³H]CP55940 (174.6 Ci·mmol⁻¹) by PerkinElmer Life Sciences Inc (Boston, MA, USA).

THCV behaves in vitro as a potent cannabinoid CB₂ receptor agonist

In our initial experiments, we investigated whether THCV shares the ability of established cannabinoid CB₂ receptor agonists (Pertwee, 1999) to inhibit forskolin-induced stimulation of cyclic AMP production in hCB₂-transfected CHO cells. As shown in Figure 2, we found that at concentrations in the nanomolar range THCV can indeed induce such inhibition in this bioassay, its EC₅₀ and E_max values with 95% CIs shown in parentheses being 38 nM (12 and 124 nM) and 40% (32 and 48%) respectively. Corresponding values for the established CB₁/CB₂ receptor agonist, CP55940, which is thought to display full (high-efficacy) agonism at the cannabinoid CB₂ receptor were 6.9 nM (3.5 and 13 nM) and 55% (50 and 60%), respectively (Figure 2).

Figure 2

The effect of THCV and CP55940 on forskolin-induced stimulation of cyclic AMP production in CHO cells transfected with hCB₂ receptors that had or had not been pre-incubated overnight with pertussis toxin (PTX; 100 ng·mL⁻¹) (n= 4). EC₅₀ …

We also obtained some confirmatory evidence that these inhibitory effects of THCV and CP55940 were cannabinoid CB₂ receptor mediated. First, as shown in Figure 2, we established that at concentrations of 0.1 nM to 10 µM, neither of these compounds affected forskolin-stimulated cyclic AMP production in CHO–hCB₂ cells when these had been pre-incubated overnight with pertussis toxin in a manner expected to eliminate G_i/o-mediated signalling (Glass and Felder, 1997; Bonhaus et al., 1998; Coutts et al., 2001). Second, we found that at such concentrations, neither THCV nor CP55940 inhibited forskolin-stimulated cyclic AMP production in untransfected CHO cells (n= 4; data not shown). Next, we showed that THCV can displace [³H]CP55940 from specific binding sites on cell membranes from CHO cells transfected with hCB₂receptors, its mean K_i value with 95% CIs shown in brackets being 225 nM (170 and 298 nM; n= 8).

We have found previously that THCV opposes CP55940-induced stimulation of [³⁵S]GTPγS binding to membranes of CHO–hCB₂ cells (Thomas et al., 2005), and that by itself THCV does not behave as a CB₂ receptor agonist in this bioassay at concentrations ranging from 0.01 nM to 10 µM (Thomas and Pertwee, unpublished). However, the CB₂ receptor density determined by [³H]CP55940 saturation binding was three times lower in the CHO cells used in these previous experiments ([³H]CP55940 B_max= 72.57 pmol·mg⁻¹) than in the cells used in the present investigation (Methods). We therefore carried out some additional experiments directed at testing the hypothesis that THCV would behave as a CB₂ receptor agonist in the [³⁵S]GTPγS assay when this was performed with membranes obtained from the CHO–hCB₂ cell line in which this cannabinoid did appear to activate CB₂ receptors (Figure 2). We found that THCV can indeed stimulate [³⁵S]GTPγS binding to membranes obtained from these CHO–hCB₂ cells. Its EC₅₀ and E_max values with 95% CIs shown in parentheses were 41.5 nM (10.5 and 164 nM) and 30% (26 and 34%), respectively (n= 6), and hence similar to the EC₅₀ and E_max values we obtained for THCV in the cyclic AMP assay. Corresponding values for CP55940 were 0.8 nM (0.3 and 2.1 nM) and 45% (40 and 50%), respectively (n= 4).

We also discovered that unlike CP55940, THCV did not inhibit forskolin-stimulated cyclic AMP production in hCB₁-transfected CHO cells, behaving instead as an hCB₁ cannabinoid receptor inverse agonist (Figure 3A). This ability of THCV to enhance forskolin-induced stimulation of cyclic AMP production was not observed in CHO–hCB₁ cells that had been pre-incubated overnight with pertussis toxin (Figure 3B). In contrast, the effect of CP55940 on cyclic AMP production switched from an inhibitory to a stimulatory effect in response to such pre-incubation with pertussis toxin (Figure 3B), presumably reflecting the reported ability of CB₁ receptors to activate G_sproteins in the absence of functional G_i/o coupling (Glass and Felder, 1997;Bonhaus et al., 1998; Howlett et al., 2002). No attempt was made to determine whether THCV-induced inhibition of forskolin-stimulated cyclic AMP production by CHO–hCB₂ cells was susceptible to antagonism by the established selective cannabinoid CB₂ receptor competitive antagonists, SR144528 and AM630. This was because, as we have also reported previously (Ross et al., 1999a,b;), both these antagonists by themselves produced marked signs of inverse agonism in the cyclic AMP assay performed with our CHO–hCB₂ cells (data not shown).

Figure 3

The effect of THCV and CP55940 on forskolin-induced stimulation of cyclic AMP production in CHO cells transfected with hCB₁ receptors that (A) had not been or (B) had been pre-incubated overnight with pertussis toxin (PTX; 100 ng·mL⁻¹ …

Our finding that THCV appears to activate human cannabinoid CB₂ receptorin vitro prompted us to investigate its ability to activate CB₂ receptors in vivoin mouse models of inflammation and inflammatory pain, there being evidence that CB₂ receptor activation constitutes a potential therapeutic strategy for ameliorating symptoms of this kind (Pertwee, 2009). Whereas human and mouse CB₁ receptors have almost identical amino acid sequences, there are significant differences between human and mouse CB₂ receptors with regard both to their primary amino acid composition and to the manner in which they respond to at least some ligands (Howlett et al., 2002; Binghamet al., 2007). Thus, for example, the selective ligand for CB₂ receptors, S-AM1241, has been reported by Bingham et al. (2007) to behave as an agonist at the hCB₂ receptor, but as an inverse agonist at mouse and rat CB₂receptors. Moreover, our evidence that THCV can activate hCB₂ receptors came from experiments with tissue obtained from a cell line in which these receptors are expressed at an extremely high level. Consequently, before proceeding to any in vivo experiments, it was first important to establish whether THCV can activate mouse CB₂ receptors in vitro in a tissue in which these receptors are expressed at physiologically relevant levels. The tissue selected for these experiments was mouse spleen as this has been reported to be well populated with CB₂ receptors (Pertwee, 1997). We found that THCV can indeed stimulate [³⁵S]GTPγS binding to these membranes. Its EC₅₀ andE_max values in these experiments with 95% CIs shown in parentheses were 69 nM (2.6 and 1804 nM) and 23% (3.8 and 42%), respectively (n= 6). Mouse spleen has been reported to express CB₁ as well as CB₂ receptors (Pertwee, 1997). Even so, it is unlikely that THCV was stimulating [³⁵S]GTPγS binding by activating CB₁ receptors in our spleen experiments because concentrations of THCV ranging from 1 to 10 µM produced no detectable stimulation of [³⁵S]GTPγS binding to spleen membranes obtained from CB₂^−/− mice (n= 6; data not shown). Moreover, THCV (n= 8; data not shown) did not share the ability of the established CB₁/CB₂ receptor agonist, CP55940, to stimulate [³⁵S]GTPγS binding to membranes obtained from mouse brain, a tissue in which CB₁ receptors are highly expressed (Howlett et al., 2002). The EC₅₀displayed by CP55940 in these experiments with its 95% CIs shown in parentheses was 10.7 nM (6.3 and 18.4 nM; n= 6).

THCV ameliorates signs of inflammation and hyperalgesia induced in mice by carrageenan or formalin

Having obtained evidence that THCV can activate naturally expressed mouse CB₂ receptors in vitro, we went on to investigate its ability to activate mouse CB₂ receptors in vivo. This we did by performing experiments directed at determining whether THCV shares the ability of established selective CB₂receptor agonists (Whiteside et al., 2007; Guindon and Hohmann, 2008) to ameliorate signs of inflammation and thermal hyperalgesia induced in rats or mice by intraplantar injection of carrageenan or formalin.

As expected, carrageenan caused the volume of each injected ipsilateral hind paw to increase relative to the volume of the contralateral uninjected paw. At 2 h after carrageenan administration, this ipsilateral hind paw oedema was significantly less in mice that had been pretreated with THCV at a dose of 0.3 mg·kg⁻¹ than in vehicle-pretreated animals (Figure 4A). Carrageenan injection also induced marked thermal hyperalgesia, as indicated by a decreased withdrawal latency of the injected paw in response to a thermal stimulus, and pretreatment with THCV at doses of either 0.3 or 1 mg·kg⁻¹ was found to reduce this thermal hypersensitivity (Figure 4B). Because carrageenan-induced oedema and thermal hyperalgesia remained at a high level throughout an observation period of 24 h (Figure 5), the anti-inflammatory and anti-nociceptive responses elicited by THCV at 0.3 mg·kg⁻¹were investigated at additional time-points. We found that the anti-oedema effect of THCV was still both present and unreduced at 3 and 4 h after carrageenan administration, slightly reduced but still present 6 h after carrageenan, and absent 24 h after carrageenan (Figure 5A). Similarly, the anti-hyperalgesic effect of THCV decreased progressively over this same observation period and was no longer detectable 24 h after carrageenan administration (Figure 5B). At no dose or time-point did THCV affect the volume or withdrawal latency of the contralateral, uninjected hind paws (data not shown).

Figure 5

Effect of THCV (0.3 mg·kg⁻¹ i.p.), administered 30 min before carrageenan (2%, 20 µL intraplantar), on (A) oedema and (B) thermal hypersensitivity evaluated at different times after carrageenan. Data represent mean values ± …

Figure 4

Effect of THCV, administered i.p. 30 min before carrageenan (2%, 20 µL intraplantar), on (A) oedema evaluated 2 h after carrageenan, and (B) thermal hypersensitivity, evaluated 3 h after carrageenan. The basal hind paw withdrawal latency displayed …

We also investigated the ability of rimonabant, a selective CB₁ receptor antagonist/inverse agonist, and SR144528, a selective CB₂ receptor antagonist/inverse agonist, to block the anti-oedema and anti-nociceptive effects of THCV. These compounds were administered 15 min before THCV (0.3 mg·kg⁻¹), and evaluations of oedema and thermal hypersensitivity were made at 2 and 3 h after carrageenan respectively. Figure 6A shows that only SR144528 was able to reverse the anti-oedema effect of THCV, suggesting that this phytocannabinoid induced its anti-inflammatory effect primarily through activation of the CB₂ receptor. However, rimonabant elicited a partial, although statistically not significant, reversal of THCV-induced anti-hyperalgesia, whereas SR144528 induced no sign of any such reversal (Figure 6B). Importantly, the doses of rimonabant (0.5 mg·kg⁻¹) and SR144528 (1 mg·kg⁻¹) used in these experiments did not affect carrageenan-induced paw oedema or thermal hypersensitivity when administered alone (data not shown). Nor did THCV at any of the doses used in this investigation affect the volume or withdrawal latency of paws that had been injected with vehicle instead of carrageenan.

Figure 6

Effect of rimonabant (RIM; 0.5 mg·kg⁻¹ i.p.) and SR144528 (1 mg·kg⁻¹ i.p.) on (A) anti-oedema and (B) anti-nociceptive effects evoked by THCV (0.3 mg·kg⁻¹ i.p.). Antagonists were administered 15 min before…

To test the ability of THCV to counteract established inflammation, the compound was administered at 0.3 mg·kg⁻¹, 30 min after carrageenan injection, and then again once daily over the next 3 days. As shown in Figure 7A,B, THCV significantly diminished oedema and thermal hyperalgesia after its first injection, and these anti-oedema and anti-hyperalgesic effects of THCV remained undiminished after each of its subsequent injections.

Figure 7

Effect of THCV (0.3 mg·kg⁻¹ i.p.), administered after carrageenan once daily for 4 days, on (A) oedema and (B) thermal hypersensitivity. Data represent mean ± SEM (n= 6–8). °°°P < 0.001 versus …

THCV also displayed dose-dependent activity against formalin-induced nociceptive behaviour (Figure 8A). More specifically, it reduced pain behaviour in both phases of the formalin test at a dose of 5 mg·kg⁻¹, decreased pain behaviour in the second but not the first phase of this test at 1 mg·kg⁻¹and did not significantly affect pain behaviour in either phase at 0.1 mg·kg⁻¹. The ameliorating effect of the higher dose of THCV (5 mg·kg⁻¹) on the first and second phases of the formalin response (Figure 8B) and of the lower dose of THCV (1 mg·kg⁻¹) on the second phase (Figure 8C) was attenuated by pretreatment both with rimonabant (0.5 mg·kg⁻¹) and with SR144528 (1 mg·kg⁻¹). The doses of rimonabant and SR144528 used in these experiments did not affect formalin-induced pain behaviour when administered alone (data not shown).

Figure 8

Effect on nociceptive behaviour evoked by formalin (1.25%, 30 µL s.c.) of THCV when administered at different doses by itself (A) or at 5 mg·kg⁻¹ (B) or 1 mg·kg⁻¹ (C) in combination with SR144528 (1 mg·kg …

This investigation was supported by grants from GW Pharmaceuticals and the National Institutes of Health (DA-03672).

Abbreviations:

AM630

6-iodopravadoline

CHO

Chinese hamster ovary

CP55940

(–)-cis-3-[2-hydroxy-4-(1,1-dimethylheptyl)phenyl]-trans-4-(3-hydroxypropyl)cyclohexanol

DMSO

dimethyl sulphoxide

PMSF

phenylmethylsulphonyl fluoride

rimonabant

N-(piperidin-1-yl)-5-(4-chlorophenyl)-1-(2,4-dichlorophenyl)-4-methyl-1H-pyrazole-3-carboxamide hydrochloride

SR144528

N-[(1S)-endo-1,3,3-trimethyl bicyclo [2.2.1]heptan-2-yl]-5-(4-chloro-3-methylphenyl)-1-(4-methylbenzyl)-pyrazole-3-carboxamide

THCV

Δ⁹-tetrahydrocannabivarin

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