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Research ArticleArticle

Differential Enantiomer-Specific Signaling of Cannabidiol at CB1 Receptors

Taryn Bosquez-Berger, Sierra Wilson, Christos Iliopoulos-Tsoutsouvas, Shan Jiang, Jim Wager-Miller, Spyros P. Nikas, Ken P. Mackie, Alexandros Makriyannis and Alex Straiker
Molecular Pharmacology December 2022, 102 (6) 259-268; DOI: https://doi.org/10.1124/molpharm.121.000305

Abstract

The two main constituents of cannabis are Δ9-tetrahydrocannabinol (THC) and cannabidiol (CBD). While Δ9-THC pharmacology has been studied extensively, CBD–long considered inactive–is now the subject of vigorous research related to epilepsy, pain, and inflammation and is popularly embraced as a virtual cure-all. However, our understanding of CBD pharmacology remains limited, although CBD inhibits cannabinoid CB1 receptor signaling, likely as a negative allosteric modulator. Cannabis synthesizes (-)-CBD, but CBD can also exist as an enantiomer, (+)-CBD. We enantioselectively synthesized both CBD enantiomers using established conditions and describe here a new, practical, and reliable, NMR-based method for confirming the enantiomeric purity of two CBD enantiomers. We also investigated the pharmacology of (+)-CBD in autaptic hippocampal neurons, a well-characterized neuronal model of endogenous cannabinoid signaling, and in CHO-K1 cells. We report the inhibition constant for displacing CP55,940 at CB1 by (+)-CBD, is 5-fold lower than (-)-CBD. We find that (+)-CBD is ∼10 times more potent at inhibiting depolarization-induced suppression of excitation (DSE), a form of endogenous cannabinoid-mediated retrograde synaptic plasticity. (+)-CBD also inhibits CB1 suppression of cAMP accumulation but with less potency, indicating that the signaling profiles of the enantiomers differ in a pathway-specific manner. In addition, we report that (+)-CBD stereoselectively and potently activates the sphingosine-1 phosphate (S1P) receptors, S1P1 and S1P3. These results provide an attractive method for synthesizing and distinguishing enantiomers of CBD and related phytocannabinoids and provide further evidence that these enantiomers have their own unique and interesting signaling properties.

SIGNIFICANCE STATEMENT Cannabidiol (CBD) is the subject of considerable scientific and popular interest, but we know little of the enantiomers of CBD. We find that the enantiomer (+)-CBD is substantially more potent inhibitor of cannabinoid CB1 receptors and that it activates sphingosine-1-phosphate receptors in an enantiomer-specific manner; we have additionally developed an improved method for the synthesis of enantiomers of CBD and related compounds.

Introduction

Cannabidiol (CBD) and Δ9-tetrahydrocannabinol (THC) are the phytocannabinoids found in greatest abundance in cannabis (Elsohly and Slade, 2005). CBD was isolated before Δ9-THC (Adams et al., 1940), but remained poorly studied for decades. Because CBD is non-euphoric and because it competes weakly with conventional orthosteric radioligands for binding at the CB1 cannabinoid receptor, (Mechoulam et al., 1970; Thomas et al., 1998), CBD has often been described as inactive. Yet different ratios of Δ9-THC and CBD in cannabis preparations yield differential effects (Hiltunen and Jarbe, 1986; Karniol and Carlini, 1973; Petitet et al., 1998; Russo and Guy, 2006), and growers have actively developed cannabis strains with different ratios of CBD and Δ9-THC satisfying a need of the commercial market. CBD is now an FDA-approved anti-epileptic (reviewed in (O'Connell et al., 2017))—although the mechanism is still uncertain—and shows promise for other potential therapeutic applications (reviewed in (Maccarrone et al., 2017)). As a consequence of this and the rapidly changing cannabis-related legal landscape, there is now a keen interest in how CBD acts in the body.

It is not widely appreciated that CBD can come in more than one form. Many chemical compounds exhibit a ‘handedness’, and CBD is no exception. Such compounds are structurally identical except that the enantiomers form mirror images of one another around a chiral center. CBD has two chiral centers and therefore has four potential stereoisomers, giving rise to two pairs of enantiomers. The cannabis plant produces one of these (R,R)-CBD, denoted here as (-)-CBD, but a series of studies of one enantiomer have reported that the non-natural (S,S)-CBD – here, (+)-CBD – may have interesting signaling properties. Several studies have reported that the non-natural CBD enantiomer (+)-CBD has a greater binding affinity when compared with its natural counterpart (Bisogno et al., 2001; Fride et al., 2004; Hanus et al., 2005). Leite et al. (Leite et al., 1982) reported that the (+)-CBD and (-)-CBD enantiomers had identical effects in a seizure model but did see a somewhat stronger effect for (+)-CBD in a pentobarbitone sleeping time test. A separate study found the enantiomers to similarly activate the capsaicin receptor TRPV1 and that (+)-CBD inhibits the uptake and subsequent hydrolysis of the endocannabinoid anandamide (AEA) (Bisogno et al., 2001). The (+)-CBD enantiomer has also been reported to possess moderate to potent anti-inflammatory properties that were both dependent and independent of the CB1 receptor (Fride et al., 2004; Hanus et al., 2005). Those studies were done at a time when it was broadly assumed that CBD was inactive at CB1 receptors, an assumption grounded in the poor ability of CBD to compete with an orthosteric agonist, such as CP55940. Although some hypothesized that CBD interferes with CB1 activation (Petitet et al., 1998), the mechanism for this was clarified only more recently by evidence for CBD as a negative allosteric modulator of CB1 (Laprairie et al., 2015). Binding at an allosteric site on CB1 would account for inhibition of CB1 signaling despite poor orthosteric binding.

Given our improved understanding of CBD activity at CB1 receptors, and the keen interest in CBD, now is a suitable time to consider how the enantiomers of CBD act at this receptor. We chose the (+)-CBD enantiomer as a starting point since previous work has focused on this enantiomer. Although the synthesis of CBD enantiomers has been previously described (Hanus et al., 2005), it remains a challenge to prove that each enantiomer, when made, is free from the other enantiomer. This is important to evaluate the biologic activity of CBD as well as THC analogs. We therefore developed a simple, reliable NMR-based method that can be applied to distinguish between the enantiomers of both CBD and THC. We then tested the signaling properties of (+)-CBD relative to (-)-CBD in two model systems. We have previously shown that (-)-CBD effectively antagonizes CB1 signaling in a neuronal model of endogenous CB1- and 2-AG-dependent plasticity (Straiker et al., 2018) and have now similarly examined (+)-CBD. We additionally tested (+)-CBD and (-)-CBD for their effects on both CB1 and sphingosine-1-phosphate (S1P) receptor signaling via the cAMP pathway. We report here that (+)-CBD signaling substantially differs from that of its natural counterpart in a pathway- and target-specific manner.

Materials and Methods

Animals and Neuronal Culture

All animal care and experimental procedures used in this study were approved by the Institutional Animal Care and Use Committee of Indiana University and conform to the Guidelines of the National Institutes of Health on the Care and Use of Animals. Mouse hippocampal neurons isolated from the CA1–CA3 region were cultured on microislands as previously described (Bekkers and Stevens, 1991; Furshpan et al., 1976). Briefly, neurons were obtained from animals (at postnatal day 0–2, killed via rapid decapitation without anesthesia) and plated onto a feeder layer of hippocampal astrocytes that had been laid down previously (Levison and McCarthy, 1991). Cultures were grown in high-glucose (20 mM) minimum essential media containing 10% horse serum, without mitotic inhibitors and used for recordings after 8 days in culture and for no more than 3 hours after removal from culture medium (Straiker and Mackie, 2005). All electrophysiological experiments were performed exclusively on excitatory neurons. All tests were made on neurons from at least two different preparations.

Electrophysiology

When a single neuron is grown on a small island of permissive substrate, it forms synapses, or ‘autapses’, onto itself. All experiments were performed on isolated autaptic neurons. Whole-cell, voltage-clamp recordings from autaptic neurons were carried out at room temperature using an Axopatch 200B amplifier (Molecular Devices, Sunnyvale, CA, USA). The extracellular solution contained (mM) NaCl 119, KCl 5, CaCl2 2, MgCl2 1, glucose 30, and HEPES 20. Continuous flow of solution through the bath chamber (∼2 mL·min−1) ensured rapid drug application and clearance. Drugs were typically prepared as a stock then diluted into extracellular solution at their final concentration and used on the same day. Recording pipettes of 1.8–3 MΩ were filled with solution containing (mM): potassium gluconate 121.5, KCl 17.5, NaCl 9, MgCl2 1, HEPES 10, EGTA 0.2, MgATP 2, and LiGTP 0.5. Access resistance was monitored, and only cells with a stable access resistance were included for data analysis.

Statistical analysis: For electrophysiology analyses, depolarization response curves were obtained to determine inhibition of excitatory synaptic transmission by endogenous 2-AG by depolarizing neurons for progressively longer durations (50 milliseconds, 100 milliseconds, 300 milliseconds, 500 milliseconds, 1 second, 3 seconds, and 10 seconds). The data were fitted with a nonlinear regression (least squares method, with top of curve (representing no effect) constrained to 1) using GraphPad Prism (La Jolla, CA). This allowed calculation of an ED50, the effective dose or duration of depolarization at which a 50% inhibition is achieved as well as the Emax. Statistically significant differences in these responses were taken as non-overlapping 95% confidence intervals (CIs). Nonsignificant differences were determined by overlapping 95% CIs and an alternative analysis of effect size. The alternative analysis was used to avoid mistakenly accepting the null hypothesis of zero difference at the 0.05 level.

Flamindo cAMP Assay

Cell culture and transfection: CHO-K1 cells were cultured in high-glucose Dulbecco’s Modified Eagle Medium/Nutrient Mixture F12 (Ham’s Medium) (Thermo Fisher Scientific, Waltham, MA, USA), while HEK293 cells were cultured in high-glucose Dulbecco’s Modified Eagle Medium (Thermo Fisher Scientific) in each case, media was supplemented with 10% fetal bovine serum and a 1% Pen/Strep solution. Cultures were maintained at 37°C with an atmosphere of 5% CO2. For the imaging experiments, the cells were dissociated using trypsin-EDTA (0.05%) and cultured on poly-D-lysine pre-coated 18-mm glass coverslips in 12-well plates. One day post-plating, the cells were transfected with the receptor of interest (rat CB1, human S1P1, or human S1P3 receptor), the fluorescent protein EYFP, and the Pink Flamindo cAMP indicator (Harada et al., 2017), using Lipofectamine 2000 Transfection Reagent (Thermo Fisher Scientific). After 3.5 hours, the transfection reagent was replaced with cell culture media and the cells used for experiments within two days of transfection.

Cell Imaging and cAMP Binding Assay

Transfected CHO-K1 or HEK293 cells, were imaged in an extracellular solution containing (mM) NaCl 119, KCl 5, CaCl2 2, MgCl2 1, glucose 30, and HEPES 20, using a Leica TCS SP5 confocal microscope with an oil-immersion 20x objective.

For experiments using CHO-K1 and HEK293 cells, the test compounds were coapplied, followed several minutes later by the adenylyl cyclase activator, forskolin (Fsk; 100 μM). Images were acquired every 30 seconds for 15 minutes and then analyzed using FIJI software with the 1-click ROI Manager plugin (Thomas and Gehrig, 2020), to measure the change in fluorescence intensity. Target cells were chosen by taking the first image in the series, increasing the brightness and marking cells that exhibited a baseline Pink Flamindo fluorescence. Occasional (< 5%) cells exhibited a high baseline fluorescence relative to the general transfected cell population. These cells were excluded from analysis since they were close to saturation. This mask of identified cells (typically 15–25 per experiment) was then applied to the image series. Baseline fluorescence intensity was normalized to 100 based on the first two minutes of the time series. Using an area under the curve (AUC) analysis for time points from 0 to 15 minutes, administration of a drug concentration series (5 nM, 50 nM, 100 nM, 250 nM, and 500 nM in CHO-K1-CB1 cells; 1 nM, 10 nM, 100 nM, 200 nM, 500 nM, and 1 μM in HEK293 wild-type cells) allowed the calculation of the IC50 for (+)-CBD in this system using Graphpad Prism 6. For a given experimental treatment, a same-day control forskolin-only experimental control was included. Experimental results were compared with their respective same-day controls using an unpaired t test.

Radioligand Binding Assay

Forebrain synaptosomal membranes were prepared from frozen rat brains by the method described by Dodd et al. (Dodd et al., 1981) and were used to assess the affinity for the CB1 binding sites using [3H]-CP55,940 (specific activity: 81.3 Ci/mmol, NDSP, NIDA) with an excess of unlabeled CP55,940 (30 mM) to determine nonspecific binding. Binding assays were performed at 37°C for 1 hour in the presence of 25 µg protein per well prior to collection of membranes by rapid filtration, washing, and scintillation facilitated detection of tritium retained on the membranes as previously described (Janero et al., 2015). The normalized data from three independent experiments were combined and analyzed using a four-parameter logistic equation to yield IC50 values that were converted to inhibition constant (Ki) values using the assumptions of Cheng and Prussoff (Cheng and Prusoff, 1973) (Table 1). Nuclear magnetic resonance (NMR) spectra were recorded in CDCl3, on a Varian INOVA-500 (1H at 500 MHz) spectrometer and chemical shifts are reported in units of δ relative to internal TMS.

View this table:
TABLE 1

Materials

Drugs: Baclofen was purchased from Sigma-Aldrich (St. Louis, MO). S1P1/S1P3 dual antagonist VPC23019, and selective S1P1 and S1P3 antagonists, W146 and TY52156 were purchased from Cayman Chemical (Ann Arbor, MI). 10mM stocks of (+)-CBD and (-)-CBD (in ethanol) were stored at -80°C and diluted shortly before use. Other drugs were initially prepared as a stock in DMSO or ethanol, then diluted using extracellular solution to their final concentration shortly before use.

Chemicals Used in Synthesis

(1R,4S)-p-mentha-2,8-dien-1-ol was purchased from AK Scientific, Union City, CA, while (+)-cis/trans-p-mentha-2,8-dien-1-ol purchased from Firmenich, Inc., Princeton, NJ. All other chemicals and solvents used in the synthesis were purchased from Sigma-Aldrich (St. Louis, MO) and included boron trifluoride diethyl etherate, anhydrous dichloromethane and pyridine, p-toluenesulfonic acid monohydrate, olivetol, and chloride (R)-3,3,3-trifluoro-2-methoxy-2-phenylpropanoyl chloride (R-MTPA).

Results

Enantioselective Synthesis and Characterization of (+)-CBD and (-)-CBD

We synthesized both the non-natural (+)-CBD [(S,S)-CBD] (2) and the natural (-)-CBD [(R,R)-CBD] (4) enantioselectively (CB1 Ki values in Table 1), following a general procedure used for the synthesis of CBD analogs (Nikas et al., 2002a; Nikas et al., 2002b; Papahatjis et al., 2002). It involves the condensation of a chiral monoterpenoid alcohol with olivetol in the presence of a catalytic amount of p-toluenesulfonic acid. Enantioselective condensation of olivetol (1) with (1R,4S)-p-mentha-2,8-dien-1-ol (purchased from AK Scientific, Union City, CA) produced (+)-CBD (2), while with (+)-cis/trans-p-mentha-2,8-dien-1-ol (purchased from Firmenich, Inc., Princeton, NJ) produced (-)-CBD (4) (Fig. 1A) (Nikas et al., 2002a; Nikas et al., 2002b; Papahatjis et al., 2002). It is well-known that (+)-CBD (2) is converted to (+)-Δ8-THC (3), after boron trifluoride etherate treatment, and similarly, (-)-CBD (4) is converted to (-)-Δ8-THC (5) (Fig. 1A) (Papahatjis et al., 2007; Papahatjis et al., 2002; Papahatjis et al., 2003; Razdan et al., 1974). Therefore, (+)-Δ8-THC (3) and (-)-Δ8-THC (5), derived from (+)-CBD (2) and (-)-CBD (4), respectively, can be used to indirectly determine the enantiomeric purity of their precursors based on the Mosher ester approach (Dale et al., 1969). Enantiomerically pure (-)-Δ8-THC (5) and (+)-Δ8-THC (3), prepared as reviewed earlier (Thakur et al., 2005) by using enantiomerically pure (-)- and (+)-verbenol (Patent: Makriyannis et al., WO2011/006099A1). Subsequently, enantiomerically pure (-)-Δ8-THC (5) and (+)-Δ8-THC (3), were derivatized with the R-MTPA chloride at the phenolic hydroxyl group to afford two diastereomers 6 (S,S,S) and 7 (R, R, S) (Fig. 1B), respectively. Comparison of the 1H-NMR chemical shifts of the methoxy groups of the diastereomeric esters showed a difference between the two diastereomers 6 and 7 (Fig. 1C). This allows the confirmation of the enantiomeric purity of both (+)-Δ8-THC and (-)-Δ8-THC. In the current study, we synthesized enantioselectively (+)-CBD (2), and we converted it to (+)-Δ8-THC (3) using identical conditions. The (+)-Δ8-THC (3) was then converted to the respective Mosher ester (6), using R-MTPA chloride. The inspection of the 1H-NMR spectrum of the resulting compound showed no cross-contamination with (-)-Δ8-THC, reflecting the enantiomeric purity of its precursor, (+)-CBD (2). Τhe same procedure was applied for the synthetic (-)-CBD to confirm that it is devoid of its enantiomer, (+)-CBD.

Fig. 1.
Fig. 1.

Enantioselective synthesis and characterization of (+)-CBD and (-)-CBD. A) Enantioselective synthesis of (+)-CBD and (-)-CBD and their transformation to (+)-Δ8-THC and (-)-Δ8-THC, respectively. B) Mosher ester derivatization of (+)-Δ8-THC and (-)-Δ8-THC. C) 1H-NMR shift of the methoxy protons of the two diastereomeric Mosher esters.

The Cannabidiol Enantiomer (+)-CBD Inhibits Neuronal CB1 Signaling More Potently Than Its Natural Counterpart

We tested whether (+)-CBD would mimic its natural counterpart’s ability to suppress synaptic transmission in autaptic hippocampal neurons (Fig. 2). Depolarization of excitatory autaptic hippocampal neurons elicits depolarization induced suppression of excitation (DSE), a form of retrograde synaptic inhibition (Straiker and Mackie, 2005). When neurons are stimulated with a series of successively longer depolarizations (50 ms, 100 ms, 300 ms, 500 ms, 1 sec, 3 seconds, 10 seconds), this results in progressively greater inhibition of neurotransmission (Straiker et al., 2011) and yields a “depolarization-response curve.” This curve permits the derivation of several pharmacological properties of endogenous cannabinoid signaling, including the calculation of an effective-dose (depolarization) 50 (ED50), the duration of depolarization that results in 50% of the maximal inhibition. A negative allosteric modulator would be expected to shift a depolarization response curve up and to the right (i.e., less DSE in a non-competitive fashion), an effect we previously observed for (-)-CBD (Straiker et al., 2018). We initially tested (+)-CBD at 1 μM based on our previous finding that (-)-CBD was fully effective at micromolar concentrations, but soon found that (+)-CBD was more potent than its counterpart. We ultimately tested (+)-CBD at 1 nM, 10 nM, 100 nM, and 1 μM, finding that 100 nM and 1 μM of CBD fully blocked DSE and that 10 nM still substantially inhibited maximal DSE responses (Fig. 3, A–B; Baseline Emax (relative excitatory postsynaptic current [EPSC] charge (95% CI)): 0.36(0.23–0.48); Emax in presence of 10 nM (+)-CBD: 0.72(0.64–0.80); 100nM (+)-CBD: 0.85(0.80–0.89); 1μM (+)-CBD: 0.65(0.49–0.80); 95% CI non-overlapping for 10 nM, 100 nM, and 1 μM versus baseline) with an IC50 of 5.5 nM (95% CI: 0.4–74). Sample DSE time courses from a single neuron treated with successively higher concentrations of (+)-CBD are shown in Fig. 3A. We confirmed our previous finding that 100 nM of (-)-CBD does not inhibit DSE (Fig. 3C, Baseline DSE in response to 3-second depolarization (± S.E.M.): 0.62 ± 0.02; DSE with 100 nM of (-)-CBD (± S.E.M.): 0.58 ± 0.04, n = 5, NS by paired t test). We confirmed that (+)-CBD alone did not inhibit EPSCs and so neither directly inhibits nor activates excitatory neurotransmission in wild-type CB1 knockout neurons (Fig. 3D, relative EPSC charge ((+)-CBD 100 nM) in WT (± S.E.M.): 1.05 ± 0.04, n = 5; in CB1 KO: 0.99 ± 0.01, n = 5).

Fig. 2.
Fig. 2.

Schematic of DSE in autaptic hippocampal neurons. A) Diagram of neuronal synapse (presynapse above) shows the steps in DSE. Post-synaptic depolarization (lightning bolt) activates diacylglycerol lipase which converts PIP2 to 2-AG. 2-AG crosses the synapse to act on presynaptic CB1 receptors and is subsequently metabolized by monoacyl glycerol lipase. CB1 activation inhibits calcium channel activation, reducing glutamate release in response to an action potential (black curved arrow). The net effect is a reduction in postsynaptic AMPA receptor activation. B) Upper panels show solitary autaptic neurons in brightfield (with attached recording pipette) and after cell filling. Lower panel shows postsynaptic currents before (green) and after inhibition from depolarization (red). C) Schematic representation of DSE shows PSCs that are repeatedly elicited by 1 ms of depolarization. These are inhibited for tens of seconds after a 3-second DSE stimulus. D) Sample time series showing EPSC responses to increasing durations of depolarization (50 ms to 10 seconds). E) Depolarization-response curve showing increased inhibition in response to longer durations of depolarization (1.0 = no effect). DSE is absent in CB1 knockout cultures.

Fig. 3.
Fig. 3.

(+)-Cannabidiol ((+)-CBD) suppresses CB1 receptor-mediated depolarization induced suppression of excitation (DSE). A) Sample DSE responses show that 10 nM but not 1 nM (+)-CBD treatment (5 minutes) inhibits CB1-mediated DSE. B) (+)-CBD shifts the response curve for DSE in a concentration-dependent manner. C) At 100 nM, (+)-CBD substantially inhibits DSE, whereas (-)-CBD does not. D) (+)-CBD (100 nM) does not directly inhibit excitatory post-synaptic currents. E) Representative time course shows that 2-AG responses (1 μM) are reversed 100 nM (+)-CBD. F) Summary results for D. G) Sample experiment showing that GABAB responses (25 μM of baclofen) are unaltered by 100 nM (+)-CBD treatment. H) Sample control time course for baclofen. I) Summary of results for G-H. Error bars indicate S.E.M.. J) S1P1/3 antagonist VPC23019 does not affect DSE. *, P < 0.05 by paired t test.

We also tested whether (+)-CBD interferes with inhibition of EPSCs by bath-applied 2-AG, allowing us to assess whether the effect of (+)-CBD was perhaps due to a post-synaptic alteration of 2-AG release. Using a sub-maximal concentration of 2-AG, 1 μM, we found that 100 nM of (+)-CBD reversed 2-AG effects (Fig. 3 E–F, relative EPSC charge 3 minutes after 1 uM of 2-AG (± S.E.M.): 0.53 ± 0.07; charge relative to original baseline with 2-AG and (+)-CBD (100 nM) (± S.E.M.): 0.84 ± 0.07, n = 5; P < 0.005, paired t test).

To rule out the possibility that the effect of (+)-CBD was due to general inhibition of presynaptic GPCR signaling, we tested for effects on inhibition of EPSCs by an agonist of GABAB, another Gi/o-coupled receptor that also presynaptically inhibits neurotransmitter release in this neuronal preparation (Straiker et al., 2002). We found that baclofen (25 µM) activation of GABAB still produced the expected inhibition in the presence of 100 nM (+)-CBD (Fig. 3, G–I, Relative EPSC charge 3 minutes after 25 μM of baclofen/100 nM of (+)-CBD (± S.E.M.): 0.40 ± 0.06, n = 3; 25μM baclofen only (± S.E.M.): 0.29 ± 0.04, n = 3). Fig. 3G shows a sample time course of a cell treated with (+)-CBD that had no response to 2-AG (1 μM) but a strong, reversible response to baclofen (25 μM), while Fig. 3H shows a separate sample time course for a cell that was treated with baclofen only. Because we found in separate experiments outlined below that (+)-CBD activates S1Px receptors, we tested whether the S1P1/S1P3 receptor antagonist VPC23019 impacts DSE, finding that it does not (Fig. 3J; baseline DSE (± S.E.M.): 0.61 ± 0.02; after VPC23019 (1 μM) (± S.E.M.): 0.58 ± 0.04, n = 5, NS by paired t test).

(+)-CBD is a Less Potent Inhibitor of CB1 Receptor cAMP Pathway Signaling

We additionally tested whether the (+)-CBD enantiomer differentially modulates CB1-mediated inhibition of cyclic AMP (cAMP) accumulation in CB1-transfected CHO-K1 cells. As a Gi/o-coupled GPCR, CB1 inhibits adenylyl cyclase and as a result disrupts cAMP formation (Howlett et al., 2002). Activity of CB1 in this pathway is measured as the inhibition of the synthesis of cAMP by adenylyl cyclase. Using Pink Flamindo, a red fluorescent protein-based cAMP indicator (Harada et al., 2017), we measured cAMP accumulation after treatment with the adenylyl cyclase activator, Fsk, in a CHO-K1 cell line transfected with the rat CB1 receptor.

We found that both (+)-CBD and (-)-CBD effectively interfered with CB1-mediated cAMP inhibition by 2-AG (2.5 μM, Fig. 4, A–B) and that there was no difference between the two enantiomers (IC50 for (+)-CBD (95% CI): 150 nM (81 nM–275 nM); (-)-CBD: 290 nM (121 nM–695 nM) overlapping 95% CIs). Using an alternative analysis of effect size, we see only a shift of 178 nM (±76 nM) from 208 nM to 386 nM. This difference is modest and is not statistically significant by a t test (P = 0.16, n = 3 per condition). (+)-CBD had no effect on cAMP accumulation in the absence of 2-AG (Fig. 4C) or CB1 (Fig. 5K) in these cells. The differing potency of (+)-CBD for cAMP signaling and suppressing DSE indicates a pathway-dependence of the effects of the enantiomers on CB1 signaling.

Fig. 4.
Fig. 4.

(+)-CBD suppresses CB1 receptor-mediated cAMP inhibition in CHO-K1 cells. A) Sample time courses in CB1 transfected CHO-K1 cells show that (+)-CBD concentration-dependently reverses 2AG-induced cAMP inhibition. B) Summarized data from multiple experiments (n = 3) and AUC analysis reveal IC50’s for the enantiomers (+)-CBD (95% CI): 150 nM (81 nM–275 nM); (-)-CBD: 290 nM (121 nM–695 nM). C) Sample time courses with CB1 transfected CHO-K1 cells show (+)-CBD has no effect on CB1 signaling in the absence of a CB1 agonist. D) Summary of (+)-CBD signaling in CB1-transfected CHO-K1 cells using AUC analysis.

Fig. 5.
Fig. 5.

(+)-CBD but not (-)-CBD is an agonist at sphingosine 1 phosphate receptors. A) Sample time courses in wild-type HEK293 cells show that (+)-CBD but not (-)-CBD reduces forskolin-induced rises in cAMP. B) Summary of (x)-CBD signaling in wild-type HEK293 cells using AUC analysis. C) Sample concentration-response time courses for (+)-CBD in wild-type HEK293 cells. D) Summarized data from multiple experiments (EC50 (95% CI): 147 nM (47 nM–460 nM)). E) Sample time course showing (+)-CBD effect in wild-type HEK293 cells. F) Sample time course showing (+)-CBD effect is abolished following pertussis toxin (PTX) treatment in wild-type HEK293 cells. G) Summary of (+)-CBD signaling in both non-PTX and PTX treated wild-type HEK293 cells using AUC analysis. H) Sample time courses showing the concentration-response for the dual S1P1/S1P3 blocker VPC23019 in wild-type HEK293 cells. I) Sample time courses show the effect of (+)-CBD is partly blocked by the S1P1 blocker W146 (100 nM) or the S1P3 blocker TY52156 (100 nM) and almost fully blocked by the dual S1P1/3 blocker VPC23019 (100 nM) in wild-type HEK293 cells. J) Summary of data with S1Px blockers using AUC analysis. K) Sample time courses show (+)-CBD had no effect in wild-type CHO-K1 cells. L) Sample time course shows the effect of (+)-CBD in S1P1-transfected CHO-K1 cells and its inhibition by S1P1 antagonist W146. M) Sample time course shows the effect of (+)-CBD in S1P3-transfected CHO-K1 cells and its inhibition by S1P3 antagonist TY52156. N) Summary of (+)-CBD signaling in CHO-K1 cells using AUC analysis. O) Sample time course shows S1P1 antagonist W146 has no effect on forskolin-induced cAMP signaling in S1P1-transfected CHO-K1 cells. P) Sample time course shows that S1P3 antagonist TY52156 has no effect on forskolin-induced cAMP signaling in S1P3-transfected CHO-K1 cells. Q) Sample time course shows that dual S1P1/3 blocker VPC23019 has no effect on forskolin-induced cAMP signaling in wild-type HEK293 cells. R) Summary of S1Px antagonist signaling in CHO-K1 and HEK293 cells using AUC analysis. *, P < 0.05 by an unpaired t test versus 1 (no effect). n = 3 for all experiments.

(+)-CBD is a Potent Agonist at Sphingosine-1 Phosphate Receptors S1P1 and S1P3

In the course of our experiments, we noted that 100 nM (+)-CBD inhibited cAMP accumulation in HEK293 cells, even in the absence of CB1 receptors (Fig. 5, A–B; inhibition of Fsk (1.0 = no effect): (+)-CBD: 0.75 ± 0.05; unpaired t test, P value < 0.05, (+)-CBD, P = 0.007, n = 3), while (-)-CBD had no effect even at a higher concentration (1 μM). Testing a range of concentrations, we found that the EC50 for (+)-CBD was 147nM (Fig. 5, C–D; EC50 (95% CI): 147 nM (47 nM–460 nM). We hypothesized that (+)-CBD might be acting on a Gi/o-coupled GPCR endogenously expressed in HEK293 cells. This was confirmed by pretreating cells with pertussis toxin (PTX) overnight, which prevented the effect of (+)-CBD in wild-type HEK293 cells (Fig. 5, E–G; inhibition of Fsk (1.0 = no effect): (+)-CBD: 0.71 ± 0.04; unpaired t test, P value< 0.05, (+)-CBD, P = 0.001, n = 3).

We have previously reported the complement of GPCRs expressed in HEK293 cells (Atwood et al., 2011). This includes several Gi/o-coupled lipid receptors that have some relationship to cannabinoid receptors, including two sphingosine-1 phosphate receptors, S1P1 and S1P3 (Selley et al., 2013). We therefore tested a dual S1P1/S1P3 antagonist VPC23019 (10 nM–1 μM), finding that it blocked the effects of (+)-CBD (100 nM) in a concentration-dependent manner (Fig. 5H). This indicates that one or both of these S1P receptors accounts for the effects of (+)-CBD. We then tested selective S1P1 and S1P3 antagonists W146 and TY52156 (each at 100 nM), finding that each partially inhibited the effects of (+)-CBD (Fig. 5, H–J; inhibition of (+)-CBD effect (1.0 = no effect): VPC23019: 0.24 ± 0.08; W146: 0.47 ± 0.07; TY52156: 0.59 ± 0.03; n = 3; unpaired t test, P value < 0.05, VPC23019, P = 0.0008; W146, P = 0.001; TY52156, P = 0.001). This suggested that (+)-CBD activates both receptors to inhibit cAMP accumulation in HEK293 cells.

To explore this further, we transfected CHO-K1 cells with either human S1P1 or S1P3 receptor. We found that (+)-CBD (100 nM) reduced cAMP accumulation in each case (Fig. 5, L–-N; inhibition of (+)-CBD effect (1.0 = no effect) S1P1: 0.62 ± 0.06; S1P3: 0.65 ± 0.05; n = 3 for each; unpaired t test, P value < 0.05, S1P1, P = 0.004; S1P3, P = 0.002), confirming that (+)-CBD is an agonist at S1P1 and S1P3 receptors. Antagonists W146 (300 nM) and TY52156 (300 nM) reversed this effect. Each antagonist (VPC23019, W146, and TY52156) was tested alone (1 μM) with Fsk to show they did not induce an effect in the absence of (+)-CBD in CHO-S1P1, CHO-S1P3, and wild-type HEK293 cells (Fig. 5, O–R).

Discussion

Cannabis sativa produces Δ9-THC and CBD as well as numerous additional phytocannabinoids. Most of these phytocannabinoids have chiral centers (Lewis et al., 2017), meaning that one or more additional enantiomers may exist: mirror images with distinct three-dimensional structures and, perhaps, different physiologic effects. CBD has two chiral centers, meaning that there are four potential stereoisomers, of which the cannabis plant selectively produces (R,R)-CBD denoted here as (-)-CBD. Several studies have examined the (S,S)-CBD enantiomer, referred to here as (+)-CBD, but these studies were done at a time when the pharmacology of CBD at CB1 receptors was still limited and also before CBD was approved as an anticonvulsant. Here we enantioselectively synthesized the two enantiomers and developed a new method to readily distinguish them. Additionally, we have tested the signaling profile of (+)-CBD relative to its natural cousin (-)-CBD. In a neuronal model of endogenous CB1 signaling, we find that (+)-CBD is an order of magnitude more potent than (-)-CBD. In contrast, the enantiomers have similar potency in the modulation of CB1 cAMP signaling in CHO-K1 cells, pointing to a pathway dependence. Interestingly, we were able to determine that (+)-CBD, but not (-)-CBD, activates two members of the cannabinoid-related sphingosine 1 phosphate receptor family, S1P1 and S1P3. This indicates that the signaling profiles of these CBD enantiomers differ substantially, both in terms of signaling pathway and receptor targets.

The NMR-based method reported here is simple, reliable, and less costly, when compared with chiral HPLC approaches, and it has been applied successfully to determine the enantiomeric purity of CBD and THC enantiomers. The method utilizes a key structural feature of the THC molecule termed “phenolic hydroxyl group”. Therefore, there is a high likelihood that this method can also be applied in other chiral terpenoid cannabinoids, including cannabis components (e.g., cannabivarin and cannabidivarin, as well as synthetic classic cannabinoids that encompass in their structure the phenolic hydroxyl group.

Because CBD is non-euphoric and competes poorly with cannabinoid receptor ligand binding to cannabinoid receptors (Thomas et al., 1998), the study of CBD pharmacology long lagged behind that of Δ9-THC. This contributed to an initial and persistent conclusion that CBD was either inactive or that any actions of CBD necessarily occurred via non-cannabinoid receptors, or perhaps even via a receptor-independent mechanism. Prior to the identification of cannabinoid receptors in the early 1990s, CBD enantiomers were employed as a tool to discern whether CBD was likely to act at a receptor or via some other means, such as alteration of lipid membrane properties. The rationale was that structurally distinct enantiomers would be expected to have differential effects on receptors. Leite et al., for instance found that (+)-CBD and (-)-CBD had identical anticonvulsive effects on a seizure model and concluded that the effect likely occurred independently of a receptor (Leite et al., 1982). A series of studies tested for enantiomer-specific effects on cannabinoid-related targets, such as endocannabinoid uptake or TRPV1 receptors (Bisogno et al., 2001), and for orthosteric binding at CB1 (Fride et al., 2004; Hanus et al., 2005). The latter reported that (+)-CBD binds more potently at the orthosteric site than (-)-CBD, a finding that we have confirmed here.

The last half-dozen years have seen a dramatic reappraisal of the role and mechanism of action for CBD. There is growing evidence that CBD acts via non-canonical cannabinoid-related receptors, such as GPR55 (Pertwee, 2008; Senn et al., 2020), and also evidence that CBD acts on CB1 receptors despite its poor competition with CB1 orthosteric ligands (Thomas et al., 2007). The likely explanation is that CBD is a negative allosteric modulator at CB1 (Laprairie et al., 2015). CBD therefore does bind to CB1 but at a secondary allosteric site that escapes observation during CP55940-based equilibrium binding assays. We therefore revisited the activity of (+)-CBD in several assays of CB1 signaling. We had previously tested the activity of (-)-CBD (Straiker et al., 2018) in excitatory autaptic hippocampal neurons, a well-characterized model of CB1-mediated plasticity. An architecturally simple system wherein a single neuron synapses onto itself, autaptic neurons express DSE, a 2-AG- and CB1-mediated form of retrograde inhibition (Straiker and Mackie, 2005). (+)-CBD proved to be ten times more potent than its cousin in this model, but (+)-CBD showed little difference in inhibiting cAMP signaling, an indicator of substantial pathway-specific differences in the signaling of these enantiomers. In principle, it is possible that the difference in (+)-CBD potency between our DSE and cAMP signaling models is due to the difference between the mouse (for DSE experiments) and rat (for cAMP experiments); we have previously reported on species differences in CB1 signaling for human versus rat in the autaptic model (Straiker et al., 2012). However, there is only a single amino acid difference between mouse and rat CB1 receptors, and this residue is not in a conserved region and has not been otherwise implicated in differential signaling between mouse and rat, nor have we noted differences in CB1 responses in autaptic cultures derived from rat (Straiker et al., 2002) versus mouse (Straiker and Mackie, 2005). We therefore consider it unlikely that a species difference between mouse and rat underlies the difference in (+)-CBD potency seen here.

Previous work has shown both the natural and non-natural CBD enantiomers may target a variety of receptors and channels (Bisogno et al., 2001; Fride et al., 2004; Hanus et al., 2005). We report here that unlike (-)-CBD, (+)-CBD activates the S1P receptor subtypes S1P1 and S1P3 with an EC50 of ∼150 nM. The S1P GPCR family consists of five subtypes, S1P1-5, receptors that, with the exception of S1P4, are expressed throughout the central nervous system (Choi and Chun, 2013; Dusaban et al., 2017; Grassi et al., 2019; Lucaciu et al., 2020). With their varying degrees of expression and function in glial cell types and neurons, S1P1 and S1P3 are implicated in neurogenesis in the developing rodent brain (Choi and Chun, 2013; Ye et al., 2016), blood brain barrier integrity (Groves et al., 2013; Spampinato et al., 2015), stress resilience (Corbett et al., 2019), neuropathic pain (Chen et al., 2019), and inflammation (Corbett et al., 2019; Dusaban et al., 2017; Spampinato et al., 2015). This has made them targets for the treatment of neurologic disorders, such as multiple sclerosis, Alzheimer’s, Parkinson’s, and Huntington’s disease (Choi and Chun, 2013; Grassi et al., 2019; Groves et al., 2013; Lucaciu et al., 2020). (+)-CBD may activate other members of this receptor family, but determining this would be outside the scope of the current study, although we hypothesize that the greater potency of (+)-CBD in our autaptic model is due to allosteric effects on DSE. While we did not see an effect of (+)-CBD on EPSCs in CB1 knockout neurons and we found the S1P1/3 antagonist VPC23019 to be without effect on DSE, it remains possible that the effects of (+)-CBD on DSE occur through some non-CB1-dependent action.

In summary, we have examined the signaling of (+)-CBD, an enantiomer of the phytocannabinoid CBD that is not produced by the cannabis plant. We find that the signaling of the (+)-CBD enantiomer differs from its cousin in a pathway- and target-dependent manner. In a neuronal model of CB1-dependent inhibition of neurotransmission, (+)-CBD proved much more potent, when compared with its effect on cAMP accumulation, relative to (-)-CBD. In addition, the (+)-CBD enantiomer was also found to activate the lipid receptors S1P1 and S1P3, which warrants further exploration into whether (+)-CBD could be used to treat or mitigate the severity of various neurologic disorders. The finding that the non-natural enantiomer is a much more potent inhibitor of CB1-dependent neuronal signaling is also noteworthy, given the therapeutic promise and widespread public use of CBD and suggests that enantiomer-specific effects of phytocannabinoids may merit further consideration and experimentation.

Acknowledgements

Imaging of the Flamindo-based cAMP sensor was made possible by the Indiana University Light Microscopy Imaging Center (LMIC).

Authorship Contributions

Participated in research design: Bosquez, Wilson, Nikas, Makriyannis, Straiker, Mackie.

Conducted experiments: Bosquez, Wilson, Jiang, Nikas, Straiker.

Contributed new reagents or analytic tools: Iliopoulos-Tsoutsouvas, Jiang, Wager-Miller, Nikas, Makriyannis, Straiker, Mackie.

Performed data analysis: Bosquez, Wilson, Nikas, Straiker.

Wrote or contributed to the writing of the manuscript: Bosquez, Wilson, Iliopoulos-Tsoutsouvas, Nikas, Makriyannis, Straiker, Mackie.

Footnotes

    • Received April 19, 2021.
    • Accepted September 5, 2022.

  • This work was supported by the National Institutes of Health National Institute on Drug Abuse [Grants DA009158, DA0141435, and DA045020].

  • I certify that any affiliations with or involvement (either competitive or amiable) in any organization or entity with a direct financial interest in the subject matter or materials discussed in the manuscript (e.g., employment, consultancies, stock ownership, honoraria, expert testimony, etc.) are noted below. All financial research or project support is identified in an acknowledgment in the manuscript Statement of Financial Interest. No author has an actual or perceived conflict of interest with the contents of this article.

  • ↵1 A.M. and A.S. contributed equally to this work.

  • dx.doi.org/10.1124/molpharm.121.000305.

Abbreviations

2-AG
2-arachidonoyl glycerol
AUC
area under the curve
CBD
cannabidiol
DSE
depolarization-induced suppression of excitation
EPSC
excitatory postsynaptic current
Fsk
forskolin
Ki
inhibition consant
MTPA
α-methoxy-α-(trifluoromethyl)phenylacetyl
NMR
nuclear magnetic resonance
THC
tetrahydrocannabinol

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CBD Enantiomer Signaling

Taryn Bosquez-Berger, Sierra Wilson, Christos Iliopoulos-Tsoutsouvas, Shan Jiang, Jim Wager-Miller, Spyros P. Nikas, Ken P. Mackie, Alexandros Makriyannis and Alex Straiker
Molecular Pharmacology December 1, 2022, 102 (6) 259-268; DOI: https://doi.org/10.1124/molpharm.121.000305
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