HICCUPS & Cannabis studies
Overview
There are numerous home remedies to get rid of hiccups, including holding your breath, drinking a glass of water quickly, having someone frighten or surprise you, using smelling salts, pulling hard on your tongue, and others.
For severe or chronic hiccups that are not cured with home treatment, medical treatments include medications, anesthesia to block the phrenic nerve, and surgical implantation of an electronic stimulator to the vagus nerve. Surgery to disable the phrenic nerve is a treatment of last resort.
The prognosis for hiccups is good. For most people, hiccups usually stop by themselves with no lingering effects. If hiccups continue, they may cause social embarrassment and distress, and chronic hiccups may result in speech, eating, and sleeping disorders.
Science & Research
Undated - Anecdotal - Hiccups by Ben.
2012 - News/ Anecdotal - Teen Says Marijuana Has Been a Lifesaver
2004 - Study - Cannabinoids Suppress Synaptic Input to Neurones of the Rat Dorsal Motor Nucleus of the Vagus Nerve
1998 - Study - Marijuana For Intractable Hiccups
1998 - News - Marijuana Cures Hiccups
I would like to submit proof for the idea that marijuana can stop hiccups.
I suffered from two attacks today. The first came on about 11:00 am. It passed after five or ten minutes.
The second arrived 6:30 pm and hung around for 2o minutes before I went to the Internet. I came across a post of yours and learned the herb stopped a case of yours.
Long and short: I went to my Vapor Brothers vaporizer and did some herb and, praise the Lord, the hiccup went away....IMMEDIATELY...practically on the first inhale!
They were starting to bother me and I'm glad they are gone. It is my wish that this helps somebody because you, Dr. Ginspoon, helped me BIG TIME!
God made cannabis, Man did not.
Man made alcohol, God did not.
- Corresponding author
B. N. Smith: Department of Cell and Molecular Biology, Tulane University, 2000 Stern Hall, 6400 Freret Street, New Orleans, LA 70118, USA. Email: bnsmith@tulane.edu
Abstract
Cannabinoids bind central type 1 receptors (CB1R) and modify autonomic functions, including feeding and anti-emetic behaviours, when administered peripherally or into the dorsal vagal complex. Western blots and immunohistochemistry indicated the expression of CB1R in the rat dorsal vagal complex, and tissue polymerase chain reaction confirmed that CB1R message was made within the region.
To identify a cellular substrate for the central autonomic effects of cannabinoids, whole-cell patch-clamp recordings were made in brainstem slices to determine the effects of CB1R activation on synaptic transmission to neurones of the dorsal motor nucleus of the vagus (DMV). A subset of these neurones was identified as gastric related after being labelled retrogradely from the stomach. The CB1R agonists WIN55,212-2 and anandamide decreased the frequency of spontaneous excitatory or inhibitory postsynaptic currents in a concentration-related fashion, an effect that persisted in the presence of tetrodotoxin.
Paired pulse ratios of electrically evoked postsynaptic currents were also increased by WIN55,212-2. The effects of WIN55,212-2 were sensitive to the selective CB1R antagonist AM251. Cannabinoid agonist effects on synaptic input originating from neurones in the nucleus tractus solitarius (NTS) were determined by evoking activity in the NTS with local glutamate application. Excitatory and inhibitory synaptic inputs arising from the NTS were attenuated by WIN55,212-2.
Our results indicate that cannabinoids inhibit transfer of synaptic information to the DMV, including that arising from the NTS, in part by acting at receptors located on presynaptic terminals contacting DMV neurones. Inhibition of synaptic input to DMV neurones is likely to contribute to the suppression of visceral motor responses by cannabinoids.
Type 1 cannabinoid receptors (CB1R) bind Delta(9)-tetrahydrocannabinol (Δ9-THC) and a number of ligands endogenously derived from the acylethanolamide lipid family (Walter et al. 2002).
The CB1R and its mRNA are found throughout the central nervous system, including the brainstem (Matsuda et al. 1993; Tsou et al. 1998; Van Sickle et al. 2001, 2003; Partosoedarso et al. 2003). In addition to their psychoactive properties, cannabinoids appear to have clinically relevant effects on autonomic output, particularly on gastrointestinal functions.
Activation of brainstem CB1Rs has anti-emetic effects (Tramer et al. 2001; Van Sickle et al. 2001, 2003), and modulates digestive motor function and coordination (Shook & Burks, 1989; Krowicki et al. 1999; Partosoedarso et al. 2003).
These effects could be induced by applying CB1R agonists directly to the dorsal vagal complex (DVC), which includes the nucleus tractus solitarius (NTS), dorsal motor nucleus of the vagus (DMV) and area postrema. Immunohistochemical results following peripheral lesions of the nodose ganglion or the vagus nerve suggested that CB1Rs might be produced and expressed within the DVC and implied that cannabinoids may act within the brainstem on local DVC circuits to modify vagal reflexes (Partosoedarso et al. 2003).
Thus, anti-emetic and gastric motor regulatory properties of cannabinoids have been identified, and the DVC appears to be a likely site of action for cannabinoid effects on autonomic functions.
In many parts of the brain, cannabinoids inhibit fast synaptic activity. After exogenous application, cannabinoid binding is thought to occur at G-protein-coupled receptors located on synaptic terminals to reduce presynaptically both GABAergic (Katona et al. 2000, 2001) and glutamatergic synaptic transmission (Shen et al. 1996; Szabo et al. 2000; Hajos et al. 2001; Morisset & Urban, 2001). Single-unit recordings from NTS neurones suggest that cannabinoids can inhibit action potential activity in some cells (Himmi et al. 1996), but the cellular mechanisms related to cannabinoid modulation of vagal motor output have not been explored.
Vagal afferent input from the gastrointestinal tract and other visceral systems terminates in the NTS. Subsequently, putative neuronal connections with preganglionic vagal motor neurones in the DMV form the basis for vagal reflex control of the upper gastrointestinal tract. Thus, activation of vagal afferents results in excitation of second order NTS neurones (Smith et al. 1998). Neurones in the NTS, in turn, exert synaptic control of vagal outflow from the DMV. This influence can be either inhibitory or excitatory, depending upon whether a glutamatergic or GABAergic interneurone is activated (Travagli et al. 1991; Davis et al. 2003). Modulation of these synapses is believed to influence visceral function profoundly (Travagli & Rogers, 2001; Browning et al. 2002; Davis et al. 2003).
Although some central actions of cannabinoids in the NTS have been identified, and application to the DVC has been shown to modify feeding and other autonomic behaviours, there is no information at the cellular level regarding the effects of cannabinoids within the DVC.
The hypothesis that CB1R activation inhibits synaptic input to DMV neurones, including gastric-related neurones, was tested using whole-cell patch-clamp recordings in brainstem slices. The presence of both the CB1R mRNA and protein in the rat DVC was confirmed, and the effects of CB1R activation on spontaneous and evoked excitatory and inhibitory postsynaptic currents (EPSCs and IPSCs) were determined in DMV neurones.
In particular, the effects of cannabinoids on input originating from the NTS were specifically examined. Modulation of this DVC circuit would be expected to have profound effects on visceral motor reflexes relevant to feeding and digestion, as well as other autonomic behaviours.
Methods
Animals and prelabelling of gastric-related neurones
Male Sprague–Dawley rats (Harlan, Indianapolis, IN, USA) 4–8 weeks of age were housed under a standard 12 h light–12 h dark cycle, with food and water provided ad libitum. All animals were treated and cared for in accordance with the rules of the Tulane University Animal Care and Use Committee and NIH guidelines.
For some experiments, a retrogradely transported viral vector that reports enhanced green fluorescent protein (EGFP) was used to identify gastric-related neurones (Jons & Mettenleiter, 1997; Smith et al. 2000; Davis et al. 2003; Glatzer et al. 2003).
Under sodium pentobarbitone anaesthesia (Nembutal, 50 mg kg−1i.p., Abbott Laboratories, Chicago, IL, USA), a laparotomy was performed and the gastric musculature was injected with an attenuated (Bartha) strain of pseudorabies virus, constructed to express EGFP (PRV-152; generously supplied by Dr L. W. Enquist, Princeton University).
Three to five injections (1 μl each) of PRV-152 at a titre of 1 × 108 to 2.4 × 108 plaque forming units (p.f.u.) ml−1 were made into the gastric wall musculature on the ventral surface of the gastric corpus using a 10 μl Hamilton syringe fitted with a 26 gauge needle.
The needle was left in place for an additional 30 s at each site before removal. A fresh aliquot of PRV-152 was thawed for each injection from frozen stock. Animals were maintained in a biosafety level 2 laboratory for up to 70 h postinjection, where they were allowed to recover. Food and water, which were monitored to ensure they were consumed at a normal rate, were provided ad libitum.
Based on previous studies of neuronal health and infection stages following inoculation of the stomach with PRV (Card et al. 1993; Rinaman et al. 1993; Davis et al. 2003; Glatzer et al. 2003), labelling in the brainstem and other areas of the brain was examined at 66–72 h postinoculation. This time period resulted in labelling sufficient to allow targeting of DMV neurones for recording, but few if any neurones were labelled in the NTS (i.e. transneuronally).
Brainstem slice preparation
Rats were deeply anaesthetized with sodium pentobarbitone (100 mg kg−1i.p.) or halothane inhalation (Sigma, St Louis, MO, USA) and killed by rapid decapitation while anaesthetized. Brains were rapidly removed and immersed in ice-cold (0–4°C), oxygenated (95% O2–5% CO2) artificial cerebrospinal fluid (ACSF) containing (mm): 124 NaCl, 3 KCl, 26 NaHCO3, 1.4 NaH2PO4, 11 glucose, 2 CaCl2 and 1.3 MgCl2, pH 7.3–7.4, with an osmolality of 295–310 mOsm kg−1. Transverse brainstem slices (350–400 μm thick) containing the caudal DVC were made using a vibrating microtome (Vibratome Series 1000; Technical Products Intl, St Louis, MO, USA), as previously described (Smith et al. 2002; Davis et al. 2003).
Slices were incubated for at least 1 h at 33–35°C in oxygenated ACSF. A single brain slice was then transferred to a submersion-style recording chamber on a fixed stage mounted under an upright microscope (BX51WI; Olympus, Melville, NY, USA) and continuously perfused with ACSF. The composition of the ACSF used for recordings was identical to that used in the dissection.
Electrophysiology
Neurones in the DMV were targeted for recording under a 40× water-immersion objective (NA = 0.8) with fluorescence and infrared-differential interference contrast (IR-DIC) optics (Olympus) using a CCD video camera, as previously described (Davis et al. 2003; Glatzer et al. 2003). Whole-cell patch-clamp recordings were made in voltage-clamp mode from DMV neurones using an Axopatch 200B or Multiclamp 700A amplifier (Axon Instruments, Union City, CA, USA).
Signals were low-pass filtered at 2–5 kHz, digitized at 88 kHz (Neurocorder, Cygnus Technology, Delaware Water Gap, PA, USA), and recorded onto videotape as well as to a PC-style computer (Digidata 1320 A, Axon Instruments).
Data were captured using the pCLAMP program suite (Axon Instruments) and analysed using pCLAMP programs or Mini-analysis (Synaptosoft, Decatur, GA, USA). Recording pipettes were pulled from borosilicate glass (Garner Glass Co., Claremont, CA, USA) and were filled with (mm): 130 K-gluconate, Cs-gluconate or KCl, 1 NaCl, 5 EGTA, 10 Hepes, 1 MgCl2, 1 CaCl2, 2.4 ATP and 3 KOH; pH 7.2 (adjusted with KOH or CsOH); biocytin (0.2%); tip resistance 3–5 MΩ. Seal resistance was typically 2–4 GΩ and series resistance, measured from brief voltage steps (10 mV, 5 ms) applied through the recording pipette, was typically <20 MΩ and was monitored periodically during the recording.
Recordings in which a >20% change in series resistance was measured during drug application were excluded from the analysis. Input conductance was estimated by measuring the current at the end of brief (20–400 ms) voltage pulses of 5–10 mV.
Current–voltage relationships were measured by stepping briefly to −120 mV and then applying 400 ms depolarizing voltage pulses at 10 mV steps in the presence of TTX. Resting membrane potential was determined by periodically monitoring the voltage at which no current was measured (i.e. removing voltage-clamp control of the neurone by switching to I= 0) during the recording.
Drug application
The CB1R agonist WIN55,212-2 (Sigma) was first dissolved in dimethylsulphoxide (DMSO) and then diluted in ACSF (final concentration of DMSO <0.1% by volume). Cyclodextran (10 mm; Sigma) was included in the ACSF as a carrier molecule to keep the agonist in solution.
Control measurements were made in vehicle; neither the DMSO nor the cyclodextran had any effect on DMV neurones at the concentrations used. Anandamide was applied as a water-soluble emulsion (Tocris, Baldwin, MO, USA). Agonists were bath applied for 5–20 min at a final concentration of 1–10 μm.
The CB1R antagonist AM251 (10 μm; Tocris) was applied for at least 10 min before application of the agonists. Chemical stimulation of neurones in the NTS was made by pressure applying l-glutamate (20 mm; 10 ms; 69 KPa) through a patch pipette (∼10 μm tip diameter) positioned at the surface of the slice (Picospritzer, Parker-Hannefin, Fairfield, NJ, USA).
The effectiveness of the glutamate in evoking action potentials was verified by applying the glutamate directly at the tip of the recording pipette to evoke unclamped, rapid voltage-dependent inward currents in the recorded neurone (i.e. the fast Na+ currents underlying action potential generation).
The pipette was then repositioned over the dorsal NTS. Slices were positioned such that ACSF flowed dorsolaterally away from the DMV to minimize any possible direct effects of glutamate on the recorded neurones during stimulation of the NTS. The GABAA receptor-linked Cl− channel blocker picrotoxin (100 μm), the glutamate AMPA/kainate receptor antagonist 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX; 10 μm), the NMDA receptor antagonist dl-5-aminophos-phonovaleric acid (AP-5; 50 μm; receptor antagonists all from Sigma) and tetrodotoxin (TTX; 1 μm; Sigma or Alomone Laboratories, Jerusalem, Israel) were added to the ACSF for some experiments.
Analysis and statistics
Effects of CB1R agonists on the frequency and amplitude of spontaneous and miniature PSCs (i.e. sEPSCs and sIPSCs) and TTX-insensitive miniature PSCs (i.e. mEPSCs and mIPSCs) were analysed within a recording using the Kolmogorov–Smirnov test, with at least 2 min of continuous activity being measured for each condition. Paired-pulse stimulation of the NTS was made using pairs of current pulses (2–150 μA, 300–400 μs, 50–100 ms pulse separation; A.M.P.I.; Jerusalem, Israel) through a concentric bipolar platinum–iridium electrode placed in the NTS (125 μm diameter; FHC, Bowdoinham, ME, USA).
Comparisons of glutamate-evoked PSC (eEPSC and eIPSC) amplitude and frequency were made for the period 5 s before and after each glutamate application. Frequency of eEPSCs and eIPSCs was defined as the difference between the number of currents in the first 5 s following glutamate application and the 5 s before the application, similar to previous analyses (Smith & Dudek, 2002; Davis et al. 2003).
Agonist-induced changes in evoked PSC frequency were determined using the ratio of the average number of evoked PSCs (minimum five glutamate stimuli) before and 10–20 min after bath application of the agonist. Pooled effects of agonist on eEPSCs and eIPSCs were assessed by averaging results obtained at 15 and 20 min intervals after agonist application.
Effects at this time point were analysed using Student's paired two-tailed t test (significance at P < 0.05), and analyses of time-dependent effects of drugs were made using ANOVA. Numbers are reported as the means ±s.e.m.
CB1R mRNA and protein expression
Animals were anaesthetized as above, the brains and spleens were removed, and sections of brainstem tissue containing the DVC (400 μm) were made using a vibratome. The DVC was then microdissected from the brainstem sections under a dissection microscope (Nikon, Melville, NY, USA) and homogenized in a lysis buffer for total RNA or protein extraction.
For total RNA extraction, sample tissues were homogenized in Ultraspec (Biotecx, Houston, TX, USA) using a microvortexer. Following centrifugation of the homogenate, the total RNA was precipitated using 0.8 v/v isopropanol, washed in 70% ethanol, and then resuspended in 100 μl of water. Reverse transcription (RT) was performed using the Superscript II first-strand synthesis system according to the manufacturer's protocol (Invitrogen, Carlsbad, CA, USA). Amplification of cDNAs via polymerase chain reaction (PCR) was performed using primers designed to amplify the CB1R (forward primer 5′-TGTGGGCAGCCTGTTCCTCA-3′; reverse primer 5′-GGGTTTTGGCCAGCCTAATGTC-3′), the CB2 receptor (forward primer 5′-CTCCTGGGCTGGCTTCTTTTCATT-3′; reverse primer 5′-CTCTCCACTCCGCAGGGCATAA-3′), and GAPDH (forward primer 5′-GGACATTGTTGCCATCAACGAC-3′; reverse primer 5′-ATGAGCCCTTCCACGATGCCAAAG-3′).
All primers used were synthesized by Integrated DNA Technologies, Inc. (IDT, Coralville, IA, USA). For these PCR reactions the following thermocycling profile was used: initial denaturation at 94°C for 5 min, 35 cycles of 94°C for 30 s, 55°C for 30 s, 72°C for 1 min, followed by a final extension at 72°C for 15 min. All RT and PCR reactions were performed using a MJ Research PTC-200 (MJ Research, Waltham, MA, USA).
For total protein extraction, sample tissues were homogenized in a lysis buffer containing: 50 mm Hepes, pH 7.4, 150 mm NaCl, 1% deoxycholate, 1% NP-40, 0.1% SDS, 1X protease inhibitor cocktail using a microvortexer. Sodium dodecyl sulphate–polyacrylamide gel electrophoresis was used to size fractionate 40 μg of each sample homogenate using 10% Tris–glycine precast gels. Following electrophoresis, the proteins were transferred to a polyvinylidene difluoride (PVDF) membrane.
Nonspecific protein binding was blocked by treating the membranes with 3% BSA in Tris-buffered saline containing 0.1% Tween-20 (TBST) for 1 h at room temperature.
The membranes were then washed with TBST buffer and the CB1R was visualized with an affinity-purified polyclonal antibody to the amide terminus of the CB1R (a gift of Dr K. Mackie, University of Washington or Alpha Diagnostic Inc., San Antonio, TX, USA; 1 h at room temperature) or the CB2 receptor (Alpha Diagnostic Inc.), followed by alkaline phosphatase conjugated to a goat anti- rabbit secondary antibody (IgG; Vector Laboratories, Burlingame, CA, USA; 30 min at room temperature) and Immunstar alkaline phosphatase chemiluminescence reagent following the manufacturer's protocol (Bio-Rad, Hercules, CA, USA)
. Receptor binding was visualized by exposing reacted blots to Kodak Biomax MR film for 2 min. Control blots were done with antibody that was pre-absorbed with the cognate peptide.
Immunohistochemistry
Immunohistochemical localization of the CB1R was performed in a similar way to previous descriptions (Tsou et al. 1998). Briefly, adult rats were anaesthetized with sodium pentobarbitone and perfused transcardially with 0.15 m sodium phosphate buffer (pH 7.4) followed by 4% paraformaldehyde in 0.15 m sodium phosphate buffer containing 0.1% picric acid. In some cases, FluoroGold (Fluorochrome, Inc., Denver, CO, USA) dissolved in 20% lactose in 0.9% saline was injected intraperitoneally (two injections separated by 15 min; 10 mg kg−1 total) 7–10 days prior to the procedure to identify the location of DMV neurones, as described by Leong & Ling (1990).
The brains were removed, postfixed for 2 h in the same fixative, immersed in 30% sucrose in 0.01 m phosphate-buffered saline (PBS, pH 7.4) until they equilibrated, and sectioned at 20–30 μm on a freezing sliding microtome. After several rinses in PBS, floating sections were immersed in CB1R antibody (Alpha Diagnostic Inc.) in PBS (1:50; 24 h at 4°C) with 0.1% Triton X-100 and 0.1% normal goat serum.
The antibody could be reused several times at this concentration. After several rinses in PBS, sections were treated with a fluorescence-conjugated (AlexaFluor 593; Molecular Probes, Eugene, OR, USA) goat anti-rabbit secondary antibody (IgG; 1:200; 2 h at room temperature), followed by more rinsing in PBS. Some sections were subsequently treated with a monoclonal antibody to synaptophysin (Chemicon, Temecula, CA, USA; 1:100; 24 h at 4°C).
The synaptophysin was visualized using a goat anti-mouse secondary antibody conjugated to AlexaFluor 488 (Molecular Probes). Sections were then mounted onto charged slides (Superfrost Plus; Fisher Scientific, Pittsburgh, PA, USA), air dried, covered in an anti-oxidant medium (Vectashield, Vector Laboratories), and coverslipped.
Sections were usually viewed under a Leica DMLB microscope and images were captured using a Spot RT digital CCD camera (Diagnostic Instruments, Sterling Heights, MI, USA). For identifying immunohistochemical colocalization of CB1R with synaptophysin, a scanning laser confocal microscope (LSM 510 META; Carl Zeiss, Thornwood, NY, USA) was used to obtain 0.36–0.72 μm optical sections.
Results
CB1R message and protein expression in the DVC
Total RNA and protein were extracted from spleen, whole brainstem, and microdissected DVC from 400 μm thick brainstem slices.
These tissue extracts were analysed by RT-PCR to determine whether CB1R message was transcribed in the DVC and by Western blot to determine whether the receptor proteins were expressed in the region. RT-PCR analysis revealed a band of approximately 424 bp in each of the tissues analysed, indicating that the CB1R was transcribed within the DVC. (Fig. 1A).
Unlike the transcripts for CB1R, CB2 receptor was not transcribed in the brainstem or DVC. However, a band of approximately 345 bp was revealed in tissue from the spleen. In the DVC, message for CBR1, but not CBR2, was actively transcribed.
Figure 1.
Expression of cannabinoid type 1 receptor (CB1R) in the dorsal vagal complex (DVC)
A, an RT-PCR product of 424 bp that corresponds to the expected size of the CB1R amplimer was expressed in samples isolated from the DVC, brainstem and spleen.
An RT-PCR product of 345 bp that corresponds to the expected size of the cannabinoid type 2 receptor (CB2) amplimer was not found in samples from the either the DVC or the brainstem, but was present in samples from the spleen.
As a positive control, a 441 bp amplimer for GAPDH was detected in each sample. B, a CB1R polyclonal antibody recognized a major band of approximately 64 kDa in samples from the DVC, brainstem and spleen.
A CB2 receptor polyclonal antibody recognized no band in samples from either the DVC or the brainstem samples, but a band of approximately 60 kDa was recognized in samples from the spleen. C, vagal motor neurones were labelled 1 week after intraperitoneal injection of FluoroGold (FG).
Arrows point to the DMV. D, in the same section as shown in C, CB1R immunoreactivity was prominent in the vagal complex. Arrows point to the DMV; arrowheads indicate examples of labelled NTS neurones; CC, central canal; AP area postrema. E, the merged image of C and D demonstrates spatial overlap of FG-labelled vagal motor neurones and CB1R.
Arrows indicate examples of double-labelled cells, which appear orange; D, dorsal; M, medial. F, confocal optical section through the DMV of a different animal (0.36 μm thickness) showing CB1R labelling (red) and synaptophysin labelling (green) in the same plane of section.
Arrows indicate examples of double-labelled puncta (orange). Scale bar is 100 μm in C–E, 30 μm in F.
Western blot analysis of CB1R revealed a major band at approximately 64 kDa in the brainstem and in the microdissected DVC tissue extracts (Fig. 1B). Also, CB1R expression was seen in total protein isolated from the spleen. This 64 kDa band is in close agreement with previous reports identifying the glycosylated form of the CB1R (Song & Howlett, 1995).
No evidence of CB2 receptor protein was found in extracts from the CNS, but a band of approximately 60 kDa was seen in total protein isolated from the spleen. Immunoneutralization of the CB1R antibody with the cognate peptide to which the antibodies were raised blocked the detection of the CB1R in brainstem, spleen and DVC tissue extracts, and immunoneutralization of the CB2 receptor antibody blocked the detection of CB2 receptor in the spleen. Thus, CB1R protein, but not CB2 receptor protein, was present in the isolated rat DVC.
To determine the spatial expression of CB1R within the rat DVC, identification of the receptors was also made immunohistochemically. Immunoreactive puncta were prominent in the NTS and DMV, and appeared to label the neuronal somata, including DMV motor neurones identified by intraperitoneal FluoroGold (Fig. 1C–E). In addition to apparent somatic labelling, CB1R was colocalized with synaptophysin in 0.36 μm confocal optical sections from the DMV (Fig. 1F).
WIN55,212-2 effects on membrane properties
The effects of the CB1R agonist WIN55,212-2 were examined to determine whether the agonist affected membrane properties of DMV neurones. Resting membrane properties (i.e. resting membrane potential, holding current and input resistance) were not overtly altered by the agonist (n= 23).
A series of voltage pulses applied in the presence of TTX resulted in a linear current–voltage relationship at potentials near the resting membrane potential, and the slope of that relationship was unaltered by WIN55,212-2 (Fig. 2; n= 9).
The rectification of the current–voltage curve observed at potentials positive to about −40 mV was decreased by the agonist in four out of nine cells. The rectification was not observed when Cs+ was used as the primary cation carrier (n= 5), suggesting that a voltage-dependent K+ current may be inhibited by WIN55,212-2 in some neurones, in the absence of an effect on resting whole-cell conductance.
Effect of WIN55,212-2 on voltage-dependent whole-cell conductance
A, Current trace showing a reduction in peak amplitude of a putative K+ current following application of 10 μm WIN55,212-2 (arrow). B, Current–voltage plot showing the outward current activated by increasing voltage steps in normal ACSF and in the presence of WIN55,212-2 (arrow). Pooled data from 4 neurones that displayed a change in rectification is shown. Inset shows the voltage protocol used to generate the current responses.
The effects on synaptic input of WIN55,212-2 were examined to determine whether overall synaptic regulation was altered in caudal DMV neurones, voltage clamped at −70 mV to examine spontaneous EPSCs (sEPSCs) and at −20 mV for spontaneous IPSCs (sIPSCs). Some neurones were labelled with EGFP after inoculation of the stomach with PRV-152 (n= 16), which allowed them to be targeted for recording and identified as gastric-related neurones (Fig. 3).
Similar to previous electrophysiological data obtained from PRV-152-infected neurones in this and other brain areas (Smith et al. 2000; Irnaten et al. 2001; Wang et al. 2001; Davis et al. 2003; Glatzer et al. 2003), the membrane properties and synaptic input patterns of these neurones did not differ from those of DMV neurones in uninfected animals.
A comparison of membrane properties of PRV-152-labelled and unlabelled DMV neurones is shown in Table 1.
The effects on synaptic input of WIN55,212-2 were examined to determine whether overall synaptic regulation was altered in caudal DMV neurones, voltage clamped at −70 mV to examine spontaneous EPSCs (sEPSCs) and at −20 mV for spontaneous IPSCs (sIPSCs).
Some neurones were labelled with EGFP after inoculation of the stomach with PRV-152 (n= 16), which allowed them to be targeted for recording and identified as gastric-related neurones (Fig. 3).
Similar to previous electrophysiological data obtained from PRV-152-infected neurones in this and other brain areas (Smith et al. 2000; Irnaten et al. 2001; Wang et al. 2001; Davis et al. 2003; Glatzer et al. 2003), the membrane properties and synaptic input patterns of these neurones did not differ from those of DMV neurones in uninfected animals. A comparison of membrane properties of PRV-152-labelled and unlabelled DMV neurones is shown in Table 1.
Identification and recording from gastric-related rat DMV neurones prelabelled with PRV-152
A, Whole-mount view of a brainstem slice (400 μm) after fixation revealed EGFP-labelled DMV neurones 62 h after inoculation of the stomach wall.
The neurones indicated (arrow and arrowhead) were each targeted for separate recordings and filled with biocytin. B, the same slice and plane of section viewed with optics demonstrating the biocytin label (i.e. avidin–rhodamine fluorescence). The filled neurones are indicated by the arrow and arrowhead.
The neurone identified with the arrowhead was slightly deeper in the slice. C, the same neurones in whole mount after being rendered opaque (Davis et al. 2003). Arrow and arrowhead indicate the two recorded neurones.
Membrane properties of unlabelled and PRV-152-labelled DMV neurones
WIN55,212-2 inhibition of sEPSC frequency. The frequency of sEPSCs in cells tested for responses to WIN55,212-2 under control conditions was 5.5 ± 0.7 Hz (range 2.6–14.4 Hz). The sEPSC frequency was decreased by WIN55,212-2 (1–10 μm) in 26 out of 30 neurones, including each of eight neurones identified with PRV-152 as being related to gastric regulation (P < 0.05; Kolmogorov–Smirnov test; Figs 4 and 5).
The effect of WIN55,212-2 was slow to develop, taking 10–20 min to reach maximum reduction in sEPSC frequency (Fig. 4E), and was seldom reversible. The inhibition was related to agonist concentration. At the highest concentration tested, WIN55,212-2 (10 μm) decreased sEPSC frequency in 13 of 14 unlabelled neurones and eight of eight identified gastric-related DMV neurones.
At this concentration, sEPSC frequency was reduced from 6.2 ± 0.9 Hz in control conditions to 3.2 ± 0.5 Hz in WIN55,212-2 at 20 min (41 ± 8% decrease; n= 22; Figs 4 and 5). Application of the ionotropic glutamate receptor antagonists AP-5 and CNQX abolished sEPSCs (n= 12; Fig. 4). The overall amplitude of sEPSCs was not significantly altered by WIN55,212-2 in these same neurones (12 ± 9% decrease; P > 0.05).
Vehicle alone had no effect on sEPSC frequency after 20–30 min application (n= 5; P > 0.05; Fig. 4E). A smaller but significant effect of WIN55,212-2 on sEPSC frequency was observed at a concentration of 2 μm in three out of of five neurones examined (29 ± 12% reduction; P < 0.05), and 1 μm WIN55,212-2 significantly decreased sEPSC frequency in one of three neurones (23 ± 15% reduction; P < 0.05; Fig. 5).
The frequency, but not the amplitude, of sEPSCs was significantly reduced by WIN55,212-2 in a concentration-related fashion in most neurones. There was no difference between effects on sEPSCs in PRV-152-labelled neurones versus unlabelled DMV cells (Fig. 5; P > 0.05).
The cannabinoid receptor agonist WIN55,212-2 inhibited spontaneous excitatory postsynaptic currents (sEPSCs) in the DMV
A, continuous recording of sEPSCs observed at a holding potential of −70 mV in control conditions. B, sEPSCs observed after 15 min bath application of 10 μm WIN55,212-2. C, the same neurone after addition of the NMDA and non-NMDA receptor antagonists, which blocked all sEPSCs.
The neurone was prelabelled with EGFP after inoculation of the gastric musculature with PRV-152.
The bottom traces in A–C are expanded portions of the boxed areas (1 s segments) of the respective top traces. D, cumulative plot of the interevent interval distribution before and after (arrow) perfusion of WIN55,212-2 in this cell (P < 0.05, Kolmogorov–Smirnov test). E, Mean time course of inhibition of the frequency of sEPSCs in the presence of 10 μm WIN55,212-2 (•; n= 22 cells). ○, responses to application of vehicle alone at identical time points after application (n= 5). * Statistical variation of the agonist responses from the frequency before drug application (P < 0.05; ANOVA).
Input resistance was unchanged by the agonist in these 22 neurones, being 308 ± 22 MΩ in control ACSF and 267 ± 22 MΩ 20 min after agonist application (P > 0.05). F, cumulative plot of interevent interval distribution before and after perfusion of WIN55,212-2 (2 μm) for a recording in the presence of TTX (1 μm; P < 0.05). G, time course of the inhibition of miniature EPSC frequency by WIN55,212-2 (2 μm) in the presence of TTX (n= 9). * Significant change from pre-agonist application in the presence of TTX.
Normalized inhibition of spontaneous postsynaptic current frequency by WIN55,212-2
The percentage inhibition of frequency of sEPSCs and sIPSCs is plotted for EGFP-labelled neurones and unlabelled neurones. The decrease in sEPSC and sIPSC frequency at three different agonist concentrations is also shown. Number of neurones is indicated above each bar.
To investigate the possible location of the receptors responsible for the decrease in sEPSC frequency, we analysed the effects of WIN55,212-2 on miniature EPSCs (mEPSCs). In the presence of TTX (1 μm) to block action potential-dependent synaptic activity, mEPSC frequency was 7.2 ± 1.7 Hz in these neurones (n= 9). Application of WIN55,212-2 (2 μm) significantly decreased the frequency of mEPSCs after 15 min in seven of these nine neurones (3.6 ± 0.5 Hz; 37.5 ± 8.3% decrease; P < 0.05; Fig. 4F and G).
No change in mEPSC frequency was observed in the remaining two neurones. No significant changes in holding current or input resistance were observed after perfusion of WIN55,212-2. Application of WIN55,212-2 most often resulted in a decrease in EPSC frequency, which persisted in the presence of TTX.
WIN55,212-2 inhibition of sIPSC frequency. The effects of WIN55,212-2 on sIPSCs were examined in neurones voltage clamped at −20 mV, using Cs+ as the primary cation carrier to block voltage-dependent K+ channels and consequently improve voltage clamp and facilitate sIPSC analysis (Smith et al. 1998; Davis et al. 2003).
Application of WIN55,212-2 (1–10 μm) reduced the frequency of sIPSCs in each of 21 DMV cells, including eight of eight identified as gastric-related motor neurones (P < 0.05; Kolmogorov–Smirnov test; Figs 5 and 6).
The effect of WIN55,212-2 was usually maximal within 15 min (Fig. 6E). In some cases, a small increase in frequency was transiently seen within 5 min of agonist application (not shown). Average sIPSC frequency for neurones in normal ACSF was 4.6 ± 1.2 Hz (range 0.4–15 Hz).
By 20 min after application of the highest concentration of WIN55,212-2 tested (10 μm), the mean frequency of sIPSCs was reduced to 1.8 ± 0.4 Hz (n= 14; 61 ± 10% reduction; P < 0.05; Fig. 6). Application of the GABAA receptor antagonist picrotoxin (n= 14) abolished sIPSCs. The amplitude of sIPSCs was not altered by WIN55,212-2 (P > 0.05). Vehicle alone had no effect on sIPSC frequency after 20–30 min application (n= 5; P > 0.05; Fig. 6E).
The effect of WIN55,212-2 was concentration related (Fig. 5). Application of 2 μm WIN55,212-2 resulted in a decrease in sIPSC frequency in each of four neurones examined (45 ± 13% reduction; P < 0.05), whereas 1 μm WIN55,212-2 decreased sIPSC frequency by 30 ± 12% (n= 4; P < 0.05). There was no difference between the effects on sIPSCs in PRV-152-labelled neurones versus unlabelled DMV cells (Fig. 5; P > 0.05).
Inhibition of sIPSCs by WIN55,212-2
A, continuous recording of sIPSCs observed at a holding potential of −20 mV in control conditions. Recording pipette contained Cs+. B, sIPSCs after 15 min bath application of 10 μm WIN55,212-2. C, the same neurone after addition of picrotoxin (100 μm), which blocked all sIPSCs.
The neurone was prelabelled with EGFP after inoculation of the gastric musculature with PRV-152.
The bottom traces in A–C are expanded portions of the boxed areas (1 s segments) of the respective top traces. D, cumulative plot of the interevent interval distribution before and after (arrow) perfusion of WIN55,212-2 in this neurone (P < 0.05, Kolmogorov–Smirnov test). E, mean time course of the inhibition of the sIPSC frequency in the presence of WIN55,212-2 (10 μm; •; n= 13).
* Statistical variation of the pooled agonist responses from the frequency before drug application (P < 0.05; ANOVA). ○, responses to application of vehicle alone at identical time points after application (n= 5).
Input resistance was unchanged by the agonist in these 13 neurones, being 345 ± 31 MΩ in control ACSF and 304 ± 38 MΩ at 20 min after agonist application (P > 0.05). F, cumulative plot of the interevent interval distribution before and after perfusion of WIN55,212 2 (2 μm) for a recording in the presence of TTX (1 μm; P < 0.05). G, time course of the inhibition of miniature IPSC frequency by WIN55,212-2 (2 μm) in the presence of TTX (n= 5).
* Statistical variation of the pooled responses from the frequency in the presence of TTX before agonist application (P < 0.05; ANOVA).
The effects of WIN55,212-2 on mIPSCs were analysed to determine whether the reduction in IPSC frequency was dependent upon action potentials in afferent neurones. In the presence of TTX (1 μm), the frequency of mIPSCs was 4.0 ± 0.8 Hz.
Bath application of WIN55,212-2 (2 μm) decreased the frequency of mIPSCs in each of six neurones tested (P < 0.05; Kolmogorov–Smirnov test; Fig. 6F and G). After 15 min application, mIPSC frequency was reduced to 1.7 ± 0.5 Hz (60 ± 16% reduction; P < 0.05). The amplitude of mIPSCs was unaltered (P > 0.05). The inhibition of IPSC frequency by WIN55,212-2 therefore did not require action potentials.
Anandamide inhibited sEPSCs and sIPSCs
In order to confirm the cannabinoid effects on synaptic frequency with another CB1R agonist and to determine the effects on isolated sEPSCs and sIPSCs, the effects of the endogenous cannabinoid receptor ligand anandamide were analysed in neurones that were voltage clamped at −70 mV (Fig. 7). In the presence of picrotoxin, which blocked GABAergic sIPSCs, the frequency of sEPSCs was 11.7 ± 2.5 Hz (8–20 Hz; n= 5). Application of anandamide (10 μm) inhibited the frequency of sEPSCs in each of these neurones (P < 0.05; Fig. 7A). The maximum effect of anandamide was reached by about 15 min, similar to that for WIN55,212-2 (Fig. 7C). Anandamide decreased sEPSC frequency to 3.5 ± 0.7 Hz (68 ± 7% reduction; P < 0.05), with no effect on sEPSC amplitude.
Inhibition of the frequency of sEPSCs and sIPSCs by the endogenous cannabinoid anandamide
A, sEPSCs observed at a holding potential of −70 mV in control conditions. Botom trace shows sEPSCs in the same neurone after bath application of 10 μm anandamide. ACSF contained picrotoxin (100 μm). B, sIPSCs observed at a holding potential of −70 mV in control conditions. Bottom trace shows sIPSCs in the same neurone after bath application of 10 μm anandamide. Recording pipette contained KCl, which made sIPSCs inward currents; CNQX (10 μm) and AP-5 (50 μm) were present extracellularly. C, mean time course of the inhibition of sEPSC frequency in the presence of 10 μm anandamide (n= 4). D, mean time course of inhibition of sIPSC frequency in the presence of 10 μm anandamide (n= 6). Asterisks in C and D indicate significant difference from predrug frequency (P < 0.05; ANOVA).
The effects of anandamide were also examined on sIPSCs at a holding potential of −70 mV (Fig. 7B). For these experiments, Cl− was used as the primary anion in the recording pipette, which reversed the polarity of GABAA receptor-mediated sIPSCs and tended to increase their amplitudes at a holding potential of −70 mV. In the presence of CNQX (10 μm) and AP-5 (50 μm) to block glutamate receptor-mediated sEPSCs, the frequency of sIPSCs was 5.6 ± 1.4 Hz (range 1.9–11.3 Hz; n= 6).
Anandamide (10 μm) reduced sIPSC frequency to 2.0 ± 0.7 Hz (57 ± 11%; P < 0.05), with no effect on sIPSC amplitude (Fig. 7B). As for sEPSCs, the maximum effect occurred by 10–15 min of application (Fig. 7D).
Blockade of CB1R
In the presence of the CB1R antagonist AM251 (10 μm) the frequency of neither sEPSCs nor sIPSCs was modulated by WIN55,212-2 (2 μm) (Fig. 8).
The frequency of sEPSCs was 2.8 ± 0.3 Hz in the presence of AM251 and 2.5 ± 0.7 Hz after bath application of WIN55,212-2 in the presence of AM251 (P > 0.05; n= 5; Fig. 8A and C).
The frequency of sIPSCs was 7.3 ± 1.6 Hz in the presence of AM251 and 6 ± 1.4 Hz after bath perfusion of WIN55,212-2 in the presence of AM251 (P > 0.05; n= 7; Fig. 8B and D).
In a few instances when AM51 was applied alone, the frequency of sEPSCs (1 of 5 cells) or sIPSCs (3 of 7 cells) was briefly increased (not shown), but returned to baseline levels within 10 min.
Although AM251 sometimes transiently increased spontaneous synaptic activity, modulation of sEPSCs and sIPSCs by WIN55,212-2 was blocked by this specific CB1R antagonist.
Inhibition of the effect of WIN55,212-2 by the CB1R antagonist AM251
A, sEPSCs recorded at a holding potential of −70 mV in the presence of a selective CB1R antagonist, AM251 (10 μm). Bottom trace shows the same neurone after 20 min bath application of WIN55,212-2 (2 μm) in the presence of AM251 (10 μm). B, sIPSCs recorded at a holding potential −20 mV (pipette contains Cs+) in the presence of AM251 (10 μm).
Bottom trace shows the same recording 20 min after addition of WIN55,212-2 (2 μm) to the bath. C, mean time course of the sEPSC frequency in the presence of WIN55,212-2 (2 μm) and AM251 (10 μm; n= 5). D, mean time course of the sIPSC frequency in the presence of WIN55,212-2 (2 μm) and AM251 (10 μm; n= 7).
Inhibition of synaptic responses evoked from the NTS
Electrical stimulation. Pairs of electrical stimuli of the NTS separated by 50–100 ms were used to generate pairs of evoked EPSCs or IPSCs (eEPSCs or eIPSCs) in DMV neurones. The effects of WIN55,212-2 were tested on responses to paired stimuli to assess further whether the CB1 agonist acted at a presynaptic site, with a change in the ratio of amplitudes suggesting a presynaptic site of action (Regehr & Stevens, 2001).
For eEPSCs evoked at a pairing frequency of 10–20 Hz, paired-pulse depression resulted in the second EPSC having a smaller amplitude than the first (Browning et al. 1999).
WIN55,212-2 reduced the amplitude of the first current to a greater degree than the second in six out of seven cells, such that the paired-pulse ratio increased from 0.7 ± 0.09 in control ACSF to 1.1 ± 0.2 in WIN55,212-2 (10 μm, n= 7; P < 0.05; Fig. 9).
The paired-pulse ratio for eIPSCs increased from 1.1 ± 0.1 in control ACSF to 1.5 ± 0.02 in WIN-55,212–2 in (10 μm, n= 7; P < 0.05; Fig. 9). The paired-pulse ratio for both eEPSCs and eIPSCs mirrored those assessed using spontaneous synaptic events and suggested an effect at receptors on presynaptic terminals of inputs that could be evoked by stimulating the NTS.
Change in paired-pulse ratio of electrically evoked EPSCs and IPSCs induced by WIN55,212-2
A, representative traces showing pairs of EPSCs evoked 100 ms apart after electrical stimulation of afferents in the NTS. B, WIN55,212-2 (10 μm) decreased the amplitude of the first response to a greater degree than the second response.
Neurone was voltage clamped at −60 mV. C, traces showing pairs of IPSCs evoked 50 ms apart after electrical stimulation of the NTS. D, WIN55,212-2 (10 μm) decreased the amplitude of both responses, with the amplitude of the first response being suppressed to a greater extent than the first. Holding potential was −10 mV.
Traces in A–D are averages of 15–20 individual responses for each condition. E, paired-pulse ratios in the presence of WIN55,212-2, normalized to the paired-pulse ratio in normal ACSF for evoked EPSCs and IPSCs. The paired-pulse ratio increase was significant for both EPSCs and IPSCs (P < 0.05).
Chemical stimulation. Since CB1R effects were observed after electrical activation of axons in the DVC and the receptors were expressed and made within the circuitry of the DVC, we tested the hypothesis that cannabinoid agonists acted upon inputs to the DMV arising from neurones in the NTS (versus fibres of passage).
To this end, local circuits were activated using l-glutamate microapplication in the adjacent NTS and WIN55,212-2 was bath applied to the tissue (Fig. 10).
Application of l-glutamate was made at the surface of the NTS (20 mm, 10 ms) and the position of the application was adjusted so that synaptic responses could easily be observed in the relative absence of any slower inward currents, which would be indicative of direct membrane responses to the glutamate.
Glutamate microapplication to the dorsal or medial NTS was effective in activating excitatory synaptic inputs to the DMV in five of 15 neurones voltage clamped at −70 mV. In these five neurones, WIN55,212-2 (2 μm) resulted in a 33 ± 3% reduction in eEPSC frequency in the 5 s following glutamate stimulation (19.8 ± 3.6 versus 13 ± 2 eEPSCs; P < 0.05; Fig. 10), but not amplitude (94 ± 7 versus 76 ± 13 pA; P > 0.05). Excitatory synaptic input to DMV neurones, which specifically arose from neuronal activity in the adjacent NTS, was inhibited by WIN55,212-2.
Attenuation by WIN-55,212-2 of synaptic input arising from the NTS
A, left panel, image of the approximate relative positions of glutamate application (black arrow) and whole-cell recording (white arrow) in the dorsal vagal complex. l-Glutamate was applied to the dorsal NTS to evoke action potentials in intact NTS neurones, which projected to recorded DMV cells.
Right panel, a diagram showing the position of the recording and stimulating pipettes relative to the slice. The putative circuit activated by the stimulus is also shown. A theoretical axon of passage without an intact soma is also indicated, but is not activated by the glutamate. B, top trace shows excitatory responses in a DMV neurone after l-glutamate application in the NTS. Application of l-glutamate (20 mm; 10 ms) to the surface of the slice resulted in a barrage of EPSCs in this DMV neurone.
Bottom trace shows that in the presence of WIN55,212-2 (2 μm, 15 min) the response to l-glutamate activation of afferent NTS neurones was attenuated. Downward arrow indicates l-glutamate application (holding potential, −70 mV). C, top trace shows that microapplication of l-glutamate to the surface of the NTS (20 mm; 10 ms) resulted in a barrage of IPSCs in this DMV neurone. Bottom trace shows that in the presence of WIN55,212-2 (2 μm, 15 min), the response to l-glutamate activation of afferent NTS neurones was attenuated. Arrow indicates glutamate application (holding potential, −20 mV; recording pipette contained Cs+).
A set of analogous experiments was conducted to determine whether IPSCs evoked by activation of NTS neurones were also modulated by cannabinoids. For these experiments, DMV neurones were voltage clamped at −20 mV, using Cs+ as the primary cation carrier. Glutamate microapplication to the NTS activated inhibitory inputs to each of five DMV neurones examined. In these cells, WIN55,212-2 (2 μm) caused a 24 ± 2% decrease in eIPSC frequency following glutamate application (from 15.8 ± 1.6 to 12 ± 1.7 Hz; P < 0.05; Fig. 10).
Inhibitory connections between the NTS and adjacent DMV were attenuated by WIN55,212-2, consistent with the hypothesis that cannabinoids modulate both excitatory and inhibitory NTS inputs to putative preganglionic vagal motor neurones in the DMV.
Discussion
Alterations in digestive function are well-known behaviours associated with activation of cannabinoid receptors. In particular, the use of Δ9-THC as an appetite stimulant and anti-emetic have been described
(Kirkham & Williams, 2001; Williams & Kirkham, 2002; Van Sickle et al. 2001). In this study, we examined the effects of cannabinoid agonists on fast synaptic input to neurones of the rat DMV, a portion of which were identified by their specific connection with the gastric musculature. It is possible that some of these neurones were actually interneurones, being labelled via synaptic contacts with DMV motor neurones, but we believe most were motor neurones for several reasons.
First, the majority of the neurones at levels of the DMV from which we recorded were motor neurones, as indicated by the FluoroGold labelling after intraperitoneal injection.
Further, we targeted neurones expressing EGFP at a postinoculation time consistent with labelling of gastric preganglionic neurones (Card et al. 1993; Rinaman et al. 1993; Davis et al. 2003; Glatzer et al. 2003), before widespread transneuronal labelling took place.
Finally, these neurones were morphologically similar to motor neurones previously described (Browning et al. 1999). Since PRV-152 selectively labelled neuronal terminal fields in the gastric musculature (i.e. versus axons of passage), the labelled neurones in the DMV were considered to be gastric related.
Previous analyses have shown that infection of central neurones with the attenuated strain of PRV used here has no significant effect on the electrophysiological properties that we studied (Smith et al. 2000;
Irnaten et al. 2001; Wang et al. 2001; Davis et al. 2003; Glatzer et al. 2003).
With increased sample size, it remains possible that PRV-152 might eventually be found to alter some properties of labelled neurones, but none of the parameters we measured in PRV-152-infected neurones were found to differ significantly from those in uninfected animals.
Variability was high for PSC frequency in both groups because of the multitude of factors contributing to this measurement, but effects of CB1 agonists were consistent.
In the parameters measured, no differences were detected in responses of labelled and unlabelled cells to CB1R agonists. Our data from gastric-related neurones does not preclude possible effects on neurones regulating other systems, but verifies that CB1R activation in the DVC includes a potent inhibition of synaptic input to neurones that specifically regulate the stomach.
We also demonstrated that CB1Rs are both transcribed and translated in neurones of the DVC in rats. Neurones in this area are likely to be associated with visceral motor regulation, especially of the gastrointestinal tract (Krowicki et al. 1999; Glatzer et al. 2003).
The results of this study provide novel evidence for a direct modulation by cannabinoid agonists of synaptic input to DMV neurones that regulate gastric activity. Application of cannabinoid agonists produced a robust inhibitory effect on both spontaneous glutamatergic and GABAergic inputs to gastric-related and unlabelled DMV neurones.
Moreover, synaptic events evoked by either electrical activation of neurones and axons in the NTS or by increasing action potential activity in NTS neurones with intact projections to the DMV were also suppressed. Activity of cannabinoids in this system supports a role for these agents in the regulation of reflexive visceral motor output at the level of the DMV in the caudal brainstem (Partosoedarso et al. 2003; Van Sickle et al. 2003).
Activation of CB1Rs
Bath application of either of the cannabinoid receptor agonists WIN55,212-2 or anandamide caused a concentration-related reduction in the frequency, but not the amplitude, of spontaneous synaptic inputs to DMV neurones.
Although effects were observed on both sEPSCs and sIPSCs, the reduction in sIPSC frequency was particularly robust. Enhancement of synaptic activity was not usually observed.
The suppression of synaptic input was blocked or attenuated by a selective CB1R antagonist, suggesting that the synaptic modulation was mediated by CB1R activation.
Previous studies have suggested that CB1R proteins or mRNA are present in this region (Matsuda et al. 1993; Tsou et al. 1998), and immunohistochemical evidence has been presented for CB1R in the DVC of ferrets (Partosoedarso et al. 2003; Van Sickle et al. 2003). We demonstrated the presence of CB1 but not CB2 receptors (Tsou et al. 1998; Chapman, 1999) in the rat DVC by Western blot.
Furthermore, immunohistochemical labelling indicated that CB1R was prominent in the DMV of rats, including on vagal motor neurones. Some of these receptors were associated spatially with synaptophysin found in neurone terminals.
These data support the hypothesis that the effects of the cannabinoid agonists were due to activation of CB1Rs in the DMV. Message for the CB1R was also present in the region, suggesting that the receptor is both transcribed and expressed within the rat DVC.
Expression of CB1R was not apparently altered by either vagotomy or nodose ganglionectomy in the ferret DVC (Partosoedarso et al. 2003), suggesting that the effects of cannabinoids in intact animals may be due to activation of receptors that are made and expressed by neurones that participate in the central aspects of the vagal reflexes.
The presence of cytoplasmic CB1R in DMV and NTS neurones in this and other studies (Van Sickle et al. 2003) may be partly due to immunolabelling of receptors that are manufactured or metabolized in DMV neurones, analogous to labelling patterns for interneurones of the hippocampus and amygdala (Katona et al. 1999, 2001).
These receptors may be expressed at the soma, as suggested by the modulation of putative voltage-activated current we observed in a minority of cells, or they might be processed in the DMV and transported to vagal neurone terminals.
Our data imply that cannabinoids act on local DVC circuits in a manner that would be consistent with modification of vagal reflexes, which is also consistent with previous reports on the effects of cannabinoids on gastric motility, the gastro-oesophageal reflex, and emesis in intact animals (Krowicki et al. 1999; Partosoedarso et al. 2003; Van Sickle et al. 2003; Hornby & Prouty, 2004).
Thus, the CB1Rs may be transcribed in NTS neurones and are present in their terminals, suggesting that cannabinoid effects on gastrointestinal vagal reflexes can occur at the level of the connection between the NTS and DMV, a mechanism which has also been proposed for the anti-emetic effects of cannabinoids (Van Sickle et al. 2003).
We found that the transfer of synaptic information from the NTS to the DMV was significantly attenuated by cannabinoid agonists. These findings are consistent with the hypothesis that activation of CB1Rs attenuates release of neurotransmitter from terminals of NTS neurones that regulate parasympathetic motor output.
Presynaptic receptors
Previous reports noted a decrease in unit activity in some NTS neurones after Δ9-THC application (Himmi et al. 1996).
In several areas of the brain and spinal cord, synaptic inputs are attenuated by activation of CB1Rs on synaptic terminals (Shen et al. 1996; Szabo et al. 2000; Katona et al. 2000, 2001; Hajos et al. 2001; Morisset & Urban, 2001; Wilson & Nicoll, 2001).
We observed a CB1R agonist-induced decrease in the amplitude of paired EPSCs and IPSCs evoked after electrical stimulation of the NTS, and the decrease in amplitude of the first pulse was greater than the decrease in amplitude of the second pulse.
The consequent increase in paired-pulse ratio suggests activity at presynaptic CB1Rs on both GABAergic and glutamatergic terminals (Regehr & Stevens, 2001; Kline et al. 2002). Inhibition of spontaneous synaptic input was also observed in the presence of TTX, which prevents action potential-dependent release of neurotransmitter.
The decrease in mIPSC and mEPSC frequency suggests that cannabinoids reduced synaptic activity by acting at a presynaptic site in the DMV.
The reduction was especially robust for mIPSCs, which is consistent with findings in a number of brain regions (see Freund et al. 2003). The slight heterogeneity of the effect on mEPSCs at the agonist concentration used may reflect different functional output of various DMV neurones.
Alternatively, receptor density, receptor location or sensitivity to CB1R agonists may be factors. It has been proposed that central CB1Rs are primarily associated with GABA terminals, whereas a lower-affinity receptor (i.e. a putative CB3 receptor) may be associated with glutamate terminals (Hajos et al. 2001; Freund et al. 2003).
The slightly increased variability of responses observed with lower concentrations of agonist is consistent with this hypothesis. WIN55,212-2 has also been reported to modulate voltage- or ligand-gated potassium channels in some neurones (Hampson et al. 2000; Schweitzer, 2000).
Similarly, we uncovered evidence for a possible effect of postsynaptic CB1R activation that could affect voltage-dependent K+ conductances regulating action potential frequency in some neurones, but changes in input resistance or holding current following the application of cannabinoid agonists in the presence of TTX were not observed in this study.
Postsynaptic alterations in membrane potential or leak conductance therefore did not appear contribute substantially to the effects of cannabinoids on synaptic input to DMV neurones. Although additional effects on the activity of cells that are synaptically connected to DMV neurones cannot be excluded, the suppression of synaptic input, in particular GABAergic input, to DMV cells by cannabinoids appears to involve activation of CB1Rs located on presynaptic terminals.
Input from the NTS
A variety of data supports the presence of an inhibitory interneurone connecting the NTS to the DMV (Raybould et al. 1989; Zhang et al. 1998).
Stimulation of the gut or oesophagus results in excitation of NTS neurones and, often, in inhibition of DMV neurones (Zhang et al. 1998).
Stimulation of the NTS evokes gastric relaxation, whereas DMV stimulation evokes gastric motor excitation (Raybould et al. 1989). Other data indicate that there may also be an excitatory connection between the NTS and DMV (Travagli et al. 1991).
To test directly whether cannabinoids can modulate synaptic input from the NTS to the DMV, l-glutamate was applied to the NTS in the slice preparation while recording evoked activity in vagal motor neurones in the presence or absence of WIN55,212-2.
Previous studies from DMV neurones in slices used electrical stimulation of the NTS region to activate inhibitory or excitatory synaptic inputs to the DMV (Travagli et al. 1991; Travagli & Rogers, 2001; Davis et al. 2003).
A major concern with this approach is the high probability of activating axons of passage in addition to intact interneuronal projections from the NTS to the DMV. This is particularly likely when stimulating the NTS, which contains abundant fibres arising from varied CNS regions that may make synaptic contact with DMV neurones.
We used electrical stimuli in the NTS to activate paired-pulse responses in DMV neurones in order to further assess the possibility that CB1Rs were located on axon terminals, but these data do not lead to the conclusion that inputs arising from the NTS were modulated by CB1R because of the coactivation of fibres of passage with this technique. l-Glutamate activates the somadendritic region of NTS cells, but axons of passage are minimally affected, if at all (Christian & Dudek, 1988; Boudaba et al. 1996; Smith & Dudek, 2002; Davis et al. 2003).
By using l-glutamate microstimulation, we were able to determine that EPSCs or IPSCs arising from intact neurones in the NTS that projected to the DMV were attenuated by WIN55,212-2. Thus, we observed a reduction in both spontaneous and evoked synaptic frequency in putative preganglionic vagal motor neurones in the presence of WIN55,212-2, which is similar to that reported in other regions of the CNS, such as substantia gelatinosa neurones in the spinal cord (Morisset & Urban, 2001), CA1 pyramidal cells of the hippocampus (Hajos et al. 2001) and substantia nigra (Szabo et al. 2000).
Relevance to gastric function
Binding of cannabinoids in the DVC would be expected to suppress putative vago-vagal reflex responses, making such a mechanism an important regulator of vagal motor neurone responsiveness.
In support of this, the gastric motor inhibition caused by intravenously administered Δ9-THC was abolished by vagotomy, and Δ9-THC applied to the dorsal surface of the medulla mimicked the effect of intravenously administered Δ9-THC (Krowicki et al. 1999).
Although neurones within the nodose ganglion of ferrets contain CB1Rs, nodose ganglionectomy does not appear to reduce the level of CB1R staining in the ipsilateral NTS (Partosoedarso et al. 2003), suggesting that the receptor is not highly expressed in the central terminals of vagal afferent neurones.
We hypothesize that cannabinoids effect second/third-order neurones in the NTS, whose activity ultimately modulates vagal motor neurone output.
Moreover, our data suggest that CB1Rs located on the terminals of these ‘premotor’ NTS neurones are a likely site of action for the central activity of cannabinoids in modulating vagally mediated effects on gastrointestinal activity. In other central systems, endogenous cannabinoids are thought to act as inhibitory retrograde signalling molecules (Katona et al. 1999; Wilson & Nicoll, 2001).
Consistent with endogenous cannabinoid activity in the DMV, we found that application of the CB1R antagonist sometimes increased baseline synaptic activation.
Whether this was in fact due to a blockade of endogenous activity, a pharmacological activation of another type of receptor, or an inverse agonist effect warrants further study.
It is reasonable to predict that endocannabinoids released from DMV neurones might provide a direct feedback inhibition of synaptic drive from the NTS when the DMV neurones are active.
Suppression of inhibitory synapses in particular would be consistent with some of the orexigenic effects of cannabinoids, while suppression of excitatory connections might be related to the anti-emetic actions of these agonists.
Alternatively, the specific motor pathway affected (e.g. cholinergic or nonadrenergic, noncholinergic) may be an important predictor of the final physiological effect of cannabinoid binding in the DMV. Currently, effects on digestive functions can only loosely be associated with specific synaptic control of DMV neurones.
It is, however, evident that cannabinoids potently suppress synaptic inputs to DMV neurones, including those that regulate gastric function.
Acknowledgements
This work was supported by National Institutes of Health Grant DK56132. Thanks to Dr K. Mackie for generously supplying the CB1R antibody, Dr D. Corey for statistical help and Drs P. Hornby and E. Partosoedarso for advice on initial experiments.
- Received April 29, 2004.
- Revision received July 20, 2004.
- Accepted July 22, 2004.
- The Physiological society 2004
References
Marijuana For Intractable Hiccups
Source: The Lancet - Volume 351, Number 9098
Authors: Ian Gilson, Mary Busalacchi
Pubdate: Sat, 24 Jan 1998
Marijuana For Intractable Hiccups
A patient with AIDS and a history of oesophageal candidosis underwent minor ambulatory surgery. He was on indinavir, and he received perioperative intravenous midazolam and dexamethasone.
The following morning he developed persistent hiccups. Chlorpromazine controlled the hiccups only during sleep. Oral nifedipine, valproate, lansoprazole, and intravenous lidocaine had no effect. Glabellar acupuncture on day six and nine terminated the hiccups for less than an hour.
Removal of a hair from the tympanic membrane on day 8 and irrigation of marcaine into the external auditory canal on day nine gave only brief relief. On day eight the patient, who had not smoked marijuana before, smoked marijuana, and his hiccups stopped.
They recurred on day nine and on day ten the patient again smoked marijuana;hiccups stopped immediately and did not recur.
On day 14 he was found to have fluconazole-resistant oesophageal candidosis on oesophagoscopy, and was treated with oral itraconazole solution and oral amphotericin B.
Intractable hiccups has been reported as an uncommon complication of AIDS; in the largest series, most cases were attributed to oesophageal candidosis and other oesophageal diseases.1 This patient did have oesophageal candidosis, but it was longstanding and his hiccups stopped before a change in treatment, so this is unlikely to be the cause of his hiccups. Midazolam2 and dexamethsone3 are the drugs most commonly associated with iatrogenic hiccups.
The patient received both shortly before the onset of hiccups, and indinavir may have prolonged the effect of midazolam by inhibiting its metabolism. Although midazolam is contraindicated in patients on protease inhibitors, it and other proscribed drugs may be inadvertently administered if the potential for drug-drug interactions is not considered.
Anecdotal reports support the use of marijuana in AIDS-related nausea and anorexia, and dronabinol is approved for treatment of AIDS wasting.
Because intractable hiccups is an uncommon condition, it is unlikely that the use of marijuana will ever be tested in a controlled clinical trial, and blinding would be difficult.
Despite federal policy which forbids the use of marijuana therapeutically, this report should be considered for hiccups refractory to other measures.
1 Albrecht H, Stellbrink HJ. Hiccups in people with AIDS. J Acquir Immun Defic Syndr 1994; 7: 735
2 de Mendonca MJT. Midazolam-induced hiccoughs. Br Dent J 1984; 157: 49
3 Vasquez JJ. Persistent hiccup as a side effect of dexamethasone treatment. Hum Exp Toxicol 1993; 13: 32.
4 Kassirer JP. Federal foolishness and marijuana. N Engl J Med 1997; 336: 366
Aurora Medical Group, Milwaukee, WI 53212, USA ( I Gilson )
Marijuana cures hiccups
MILWAUKEE, WISCONSIN. Two American medical doctors report the use of marijuana to cure persistent (intractable) hiccups in an AIDS patient.
The patient had received intravenous midazolam and dexamethasone prior to a minor surgical operation. Following surgery he developed intractable hiccups. Chlorpromazine controlled the condition only during sleep and nifedipine, valproate, lansoprazole, and intravenous lidocaine had no effect.
Eight days after surgery the patient who had not previously smoked marijuana did so with the result that the hiccups stopped. They recurred the next day, but disappeared permanently on the next day when he again smoked marijuana. The doctors conclude that marijuana may be effective in stopping hiccups untreatable by other means.
Gilson, Ian and Busalacchi, Mary. Marijuana for intractable hiccups. The Lancet, Vol. 351, January 24, 1998, p. 267 (research letter)
An attack seizes Chaz Moore’s body, stealing much of his breath. Spasms in his throat, lungs and diaphragm cause the 17-year-old to speak in hiccups, one syllable at a time.
He says it feels like a grown man is jumping on his chest as the muscles in his belly roll like waves.
Chaz opens a jar labeled MMJ, pulls out some fresh green buds and crumbles the marijuana into a small pipe. He lights up the bowl and inhales as deeply as possible through the spasms, turning to blow smoke out his bedroom window.
A second puff, a deep cough and the attack passes.
Chaz is one of 41 people younger than 18 in Colorado who have a medical marijuana license, according to the most recent data available from the Colorado Department of Public Health and Environment.
And he’s convinced that marijuana is saving his life.
His doctors have told him he is one of about 50 people in the world diagnosed with myoclonus diaphragmatic flutter, an affliction causing muscle spasms that can recur dozens of times a day.
Until a couple of years ago, Chaz was a healthy kid, except for childhood asthma that he was outgrowing. He played in his school band and on a baseball team.
Then he started getting hives and the mysterious spasms. At first, the attacks came three to five times a week and his family rushed him to the hospital each time. Doctors tried treating him for allergies and gave him inhalers, along with high doses of painkillers and anti-anxiety drugs to relax his body.
“One week, we went nine times to the ER,” says his dad, Shan Moore. “We were going nuts, just totally freaking out. Nobody knew what was wrong.”
Doctors in Colorado Springs referred the family to National Jewish Health in Denver, where Chaz had an attack in an exam room.
One of the doctors who observed the spasms had treated a patient with the same rare illness nearly 20 years ago.
Chaz finally had a diagnosis and began treatment at Children’s Hospital Colorado, where his pediatric neurologist tried a variety of medications. At one point, he was taking a cocktail of 16 pills three times a day.
The medications would work for a time but not consistently.
So Shan Moore says he raised an “insane” idea with Chaz’s doctor — marijuana. He had seen reports online that it might help patients with multiple sclerosis.
The father says he hesitated to consider marijuana in part because his relationship with the drug goes “way back.”
Shan Moore first tried marijuana at age 10, became a self-described pothead and used everything else he could get his hands on. By his mid-30s, he says, he was dealing drugs and wound up in prison for three years when Chaz was 7.
Now 41, Moore says he has been clean for several years. The last thing he wanted to consider was getting his son started on marijuana.
But the effects of the high doses of the prescription drugs were also alarming. The family decided to give marijuana a try.
Chaz says he had tried pot once before and didn’t like feeling high.
Now he rarely experiences that feeling because the family shops for low-potency marijuana.
He has fine-tuned his medication during the past year. He starts each day with edibles such as marijuana-infused peanut butter and jelly sandwiches or marijuana cheesecake.
The food has higher levels of chemicals that seem to fend off Chaz’s attacks and stay in his system longer, without the psychoactive effects that cause a high. When an attack strikes, he smokes for immediate relief.
His friends have never hit him up for marijuana, Chaz says, and he believes kids who abuse the drug are harming patients.
“You’re taking away from my medicine,” he says. “Even though you’re out there enjoying it, you’re messing with my medicine.”
Chaz no longer uses other medications, but the marijuana created a problem.
His school district, Harrison School District 2, refused to allow the school nurse to give him marijuana. The family switched Chaz to a closer high school, hoping he could walk home when he had an attack, use marijuana and then walk back.
But the family says district officials didn’t like that idea, either, telling them they feared Chaz would be impaired and disruptive.
District spokeswoman Jennifer Sprague declined to discuss Chaz’s case and says federal and state law bar the district from administering marijuana.
“I was doing fine,” Chaz says. “I wasn’t disrupting anybody. My eyes weren’t red. I wasn’t smelling of pot. I was doing all of my work and wasn’t hurting anyone.”
Last April, after he started having as many as 35 attacks a day, Chaz enrolled in an online school.
Now he says he feels like he’s on house arrest, stuck in his bedroom with a small Dell laptop.
He’s lonely and says he sometimes loses track of what day it is because of the monotony. He’s more than a year behind his peers but determined to get an education and become a counselor for kids in hospitals.
His dad shakes his head over the battles they’ve fought.
“Medical marijuana saved his life but ruined it all at the same time,” Shan Moore says.
The family spends about $1,000 a month on various marijuana products and shops at five Colorado Springs dispensaries. The father and son have become regulars on the pro-marijuana circuit, speaking at conventions.
Being so vocal about the benefits of marijuana has been costly. The father says he lost one job because his bosses didn’t like having such an outspoken employee. He now splits wood and trims trees, picking up jobs where he can. His wife works at a Denny’s.
Chaz is on Medicaid.
The father says, altogether, they visited emergency rooms 117 times before starting marijuana. Now Chaz hasn’t been to the ER for more than a year and only goes to the doctor for routine checkups.
He doesn’t like marijuana — the taste of the food or the smell of the smoke. He feels guilty using it in the home he shares with his grandmother. He knows the damage drugs can do to a family. Right now, he sees no other options.
“If I couldn’t access marijuana,” Chaz says, “I would probably be dead.”
