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TOXICOLOGY
Departments of Anesthesiology (W.K.), Applied Physiology (K.J.-R., F.L.-H.), and Forensic Medicine (A.A.), Ulm University, Ulm, Germany; Department of Biochemistry, University College Cork, Cork, Ireland (J.J.A.H., G.O.); and Department of Neurology, Hannover University, Hannover, Germany (F.S., J.B.)
Received March 21, 2005; accepted June 2, 2005.
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Abstract |
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-bungarotoxin, a specific antagonist of the nicotinic acetylcholine receptor (nAChR), abolished these MDMA effects. The nAChR agonistic action of MDMA was confirmed by patch-clamp measurements of ion currents on human embryonic kidney cells expressing nAChR. We conclude that the neuromuscular junction is a target of MDMA and that an activation of nAChR contributes to the muscle-related symptoms of MDMA users. The drug may be of particular risk in individuals with abundant extrajunctional nAChR such as in generalized denervation or muscle regeneration processes and may act on central nAChR.
). Furthermore, MDMA activates monoamine transporters and inhibits serotonin reuptake. Besides its affinity to serotonin recognition sites (e.g., reuptake sites), MDMA exhibits dopaminergic, cholinergic, histaminergic, and adrenergic effects (Battaglia and De Souza, 1989; Mechan et al., 2002).
The most common side effects of MDMA include fasciculations, muscle pain, muscle cramps, and trismus (Henry, 1992; Nimmo et al., 1993). More severe complications such as rapidly progressing hyperthermia, hyperkalemia, metabolic acidosis, excessive creatine kinase (CK) elevation, and rhabdomyolysis have been reported rarely but even at low MDMA dosages (Screaton et al., 1992; Hall et al., 1996). These adverse effects point to the skeletal muscle or the neuromuscular junction as a target of MDMA outside the central nervous system. Metabolic myopathies have been postulated to be the cause of these severe crises (Henry et al., 1992), and recently, MDMA has been proposed to uncouple oxidative phosphorylation in skeletal muscle mitochondria by an indirect mechanism (Rusyniak et al., 2005). Toxic rhabdomyolyses have been reported in healthy individuals for a variety of substances, including statins, amphetamines, opiates, and succinylcholine (SCh). Despite different initiation, muscle breakdown finally is linked to membrane destabilization and ATP depletion, leading to cellular hypoxia, rhabdomyolysis, and edema. Muscle swelling in the tight envelope of fascia can damage the nerves and blood vessels by exerting direct pressure, i.e., cause a compartment syndrome (Martyn et al., 1992; Breucking et al., 2000; Evans and Rees, 2002). In genetically disposed swine, MDMA has been reported to trigger malignant hyperthermia (MH) in vivo (Fiege et al., 2003).
The purpose of our study was to determine whether MDMA can cause some of its adverse effects peripherally via the skeletal muscle. Therefore, we tested whether MDMA increases cytoplasmic [Ca2+] and triggers contractures in isolated skeletal muscle of normal and MHS individuals. When we identified that MDMA and SCh had similar effects on muscle and that the effects were blocked by -bungarotoxin, we extended the study in a previously unexpected direction.
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Materials and Methods |
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MDMA Analysis by Gas Chromatography (GC)/Mass Spectrometry. Centrifuged serum samples were mixed with deuterized MDMA (d5-MDMA) as internal standard before solid-phase extraction (HX 11; Separtis, Grenzach-Wyhlen, Germany). Homogenized ecstasy pills were extracted with methanol. Both types of extracts were derivatized with trifluoraceticanhydride (TFA) and evaporized (60°C; 30 min). The residues were dissolved in ethyl acetate for GC/mass spectrometry analysis performed on a Hewlett Packard 5890 GC interfaced with a Hewlett Packard 5971 mass selective detector. The carrier gas was helium. Injector and detector temperatures were 250 and 280°C, respectively. Oven temperature was set at 100°C, held for 2 min, and then increased to 280°C at 15°C/min and maintained for 1 min. The total run time was 15 min. Data were acquired with a Hewlett Packard G1034C MS ChemStation. The TFA derivatives were identified with the following masses: 135, 154, and 162 for TFA-MDMA; and 158 and 164 for TFA-d5-MDMA. In addition, caffeine was qualitatively determined.
Contracture Measurements. The IVCT was performed according to the European protocol (Ørding et al., 1997). This test determines the sensitivity of fresh muscle strips separately exposed to caffeine and halothane at cumulative concentrations in an organ bath containing Krebs-Ringer solution (118 mM NaCl, 3.4 mM KCl, 0.8 mM MgSO4, 1.2 mM KH2PO4, 11.1 mM glucose, 25.0 mM NaHCO3, and 2.5 mM CaCl2, pH 7.4). Muscle strip force was measured with a mechanoelectrical transducer (FT03; Grass Instruments, Quincy, MA). An increase in force by active shortening induced by drugs was considered as a contracture if a threshold value of 
2 mN was reached. According to the protocol, contractures were considered as pathological if they occurred at concentrations of 
2 mM caffeine or 2% halothane. Patients with pathological contractures to both substances were classified as MHS, individuals whose strips pathologically reacted to only one test reagent were classified as MH-equivocal, and the absence of pathological contractures was classified as MH-negative (MHN). Muscle strips were considered as viable if their twitch amplitudes to supramaximal electrical stimulation (pulse frequency, 0.2 Hz; pulse width, 1 ms) were >10 mN.
The effects of MDMA and SCh were also tested on muscle strips, either alone or in combination with caffeine or halothane. In this case, MDMA or SCh were added to the organ baths 10 min before the caffeine/halothane challenge. Shifts in the dose-response curves for caffeine or halothane in the presence of MDMA or SCh were determined as the difference between the lowest concentration (e.g., with and without MDMA) at which contractures of 2 mN occurred. This is in contrast to the usual chemical procedure at which concentrations leading to half-maximum contractures are determined and subtracted from each other. The reason for the different protocol is the experience that application of concentrations that usually cause maximum contractures often results in rupture at the tied ends of the muscle fiber segments. The disadvantage of the European protocol is that a shift of the steepness of the dose-response relationship is not taken into account for determination. Here, we compensate for this lack by comparing contracture amplitudes as shown in Fig. 3B.
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; Carvalho et al., 2002; Rusyniak et al., 2005), we used 100 to 1000 (4000) µM.
Mutation Analysis. For mutational screening, MH susceptibility hot-spot exons 17, 38, 39, 40, and 45 of the gene encoding the ryanodine receptor type 1 of skeletal muscle (RyR1), the Ca2+ release channel of the SR, were directly sequenced using primers as described previously (Klingler et al., 2002).
Cell Culture. Human muscle samples (50–200 mg) were mechanically teased and treated for 1 h at 37°C with collagenase (330 U/ml) (C6885; Sigma Chemie) and dissolved in Ham's F-12 medium (F0815; Biochrom, Berlin, Germany). The resulting suspension was filtered through a 20-µm nylon mesh. Myoblasts were seeded on poly-L-ornithine-coated glass coverslips. Cells were first kept in growth medium containing 5% fetal calf serum (C-23060; PromoCell, Heidelberg, Germany) at 37°C and 5% CO2. After 4 to 5 days, cell differentiation was induced by reducing serum content. Within 1 week, the myoblasts became confluent and started to fuse. Myotubes were identified by their polynucleated appearance.
Ca2+ Fluorometry. Changes of intracellular [Ca2+] were measured by use of fura-2. For dye loading, the myotubes were incubated for 30 min with 2 µM fura-2 acetylmethyl ester (Calbiochem, Bad Soden, Germany) in standard external solution (140 mM NaCl, 2.7 mM KCl, 1.0 mM MgCl2, 6.0 mM glucose, 1.5 mM CaCl2, and 12.0 mM HEPES, pH 7.3) at 37°C. Uptake was facilitated by addition of 0.02% Pluronic acid (F-127; Sigma Chemie). For the relative calibration of Ca2+ signals, the fluorescence ratio during alternating excitation at 340 and 380 nm was obtained.
MDMA was administered to the bath using a superfusion system (L/M-SPS-8; List Electronics, Darmstadt, Germany) that permitted drug application to a single cell or a small group of cells in a highly reproducible manner. Selection among the eight supply vessels connected to the multibarreled inlet pipette was controlled with magnetic valves. Pressure to the supply vessels was adjusted using a multipressure control unit (MPCU-3; Lorenz, Lindau, Germany). All Ca2+ fluorometry measurements were performed at room temperature (20°C).
Measurements on Isolated SR. Heavy SR was prepared from hind limb muscles of five sacrificed rats by homogenization and differential centrifugation as reported previously (O'Sullivan et al., 2001). The final SR pellet was resuspended in 0.1 M KCl solution and cryopreserved in liquid nitrogen. For spectrophotometry, the isolated SR was incubated with the Ca2+ chelometric dye antipyralazo III in a total volume of 2 ml using a ground glass-stoppered glass cuvette and a medium containing 19 mM MOPS, 93 mM KCl, 7.5 mM sodium pyrophosphate, 1 mM MgATP, 5 mM creatine phosphate, 20 ml of CK, and 250 mM antipyralazo III at pH 7.0. Ca2+ flux was monitored continuously with a Hewlett Packard 8452A diode-array spectrophotometer operating in dual wavelength mode at 710 and 790 nm at 37°C and constant cuvette stirring. The rate of Ca2+ uptake was calculated from the first seven 20-nmol pulses of Ca2+ added to the cuvette with a Hamilton dispensing microliter syringe. When the SR was maximally loaded with Ca2+, usually after 15 pulses, putative releasing agents were added at varying concentrations to establish whether Ca2+ release occurred. Halothane was added from a concentrated stock solution made up in pure ethanol. SR protein concentration was measured as reported previously (O'Sullivan et al., 2001).
Proton Release Measurements. The metabolism of cultured myotubes was monitored using a pH-sensitive microphysiometer (Molecular Devices, San Diego, CA) as described previously (Klingler et al., 2002). Briefly, the cells in the measuring chamber were superfused (37°C; 50 µl/min) using a computer-controlled rolling pump that alternately was switched on and off. The medium (F-12 medium; N6760; Sigma Chemie, with equimolar substitution of sodium bicarbonate by NaCl, total osmolarity 290 mOsM, pH 7.35) was only weakly buffered. During the pump-on interval the proton concentration in the chamber equilibrated with that of the medium. After stopping the perfusion, the biosensor voltage decreased (i.e., pH decreased) because protons that were extruded by the cells accumulated in the chamber until the pump was switched on again. The slope of the pH decrease gives the acidification rate corresponding to the metabolic activation of the myotubes. Pump cycle control, data acquisition, and rate calculation were carried out with the system's microcomputer (Macintosh Power PC 7600/132) and the Cytosoft program supplied with it (Molecular Devices, San Diego, CA).
Whole-Cell Patch-Clamp Recordings. Transformed human embryonic kidney (HEK) 293 cells were transfected with cDNA of mouse , 
, 
, and 
nAChR subunits. Patch-clamp measurements were performed on small cells lifted from the bottom for rapid application experiments using standard methods. The patch pipettes contained 140 mM KCl, 11 mM EGTA, 10 mM HEPES, 10 mM glucose, and 2 mM MgCl2. HEK293 cells were superfused with an extracellular solution containing 162 mM NaCl, 5.3 mM KCl, 2 mM CaCl2, 0.67 mM NaH2PO4, 0.22 mM KH2PO4, 15 mM HEPES, and 5.6 mM glucose. The pH of both solutions was adjusted to 7.3. MDMA and pancuronium were obtained from Sigma Chemie. Data were recorded with an Axopatch 200B patch-clamp amplifier (Axon Instruments Inc., Union City, CA). Membrane currents were sampled with 20 kHz using a Digidata 1200 Interface and the pCLAMP6 software suite on a PC (Axon Instruments Inc.). Data were filtered at 5 kHz for further analysis. The holding potential of the cells was kept at -40 mV. A piezo-driven, double-barreled ultrafast perfusion system was used for application of the agents to excised outside-out membrane patches or small cells. The time for solution exchange was regularly <100 µs (Krampfl et al., 2002). For the quantitative evaluation, four to 12 current traces were averaged for each experiment.
Statistical Analysis. Values showing normal distribution are presented as means and S.D. if not indicated otherwise. Values without normal distribution and discrete data are given as median, and 95% percentiles as indicated. The significance of differences between groups was evaluated by use of Wilcoxon matched pairs signed rank tests. P values less than 0.05 are considered significant.
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Results |
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Muscle Strip Contractures Induced or Potentiated by MDMA and SCh. A first set of experiments tested the potency of cumulative MDMA to elicit muscle contractures in vitro. Muscle strips from three of eight MHS patients reacted to 500 µM MDMA: strips of the R614C carrier contracted at 500 µM (Fig. 1A); strips of two patients with unknown RyR1 mutations at 1 and 2 mM, respectively; and the strips of the MH-equivocal patient at 2 mM MDMA (not shown). Also, eight MHN muscle strips were exposed to stepwise increasing concentrations of MDMA. None of the eight MHN individuals exhibited contractures even when the MDMA concentration reached 4 mM. Cumulative SCh up to 8 mM did not elicit contractures in MHS or in MHN muscle strips.
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Intracellular Ca2+ Level in Myotubes. Muscle contractures result from a myoplasmic [Ca2+] exceeding the mechanical threshold. Caffeine and halothane are known to raise myoplasmic [Ca2+] (Herrmann-Frank et al., 1999). This increase can be determined by measuring fluorescence changes of Ca2+-specific dyes in myotubes. Transient MDMA stimulation resulted in all 40 myotubes in a phasic Ca2+ signal characterized by a rapid increase, dose-dependent peak values and slow return to the resting level (Fig. 4). In an additional 10 myotubes, the nAChR was blocked with -Bgt (0.1 µM; 15 min), leading to a nondetectable amplitude or at least to a drastically reduced signal as in Fig. 4.
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Metabolic Activation of Myotubes by MDMA. An enhanced cell metabolism, e.g., originating from an increased myoplasmic [Ca2+], can be studied by measuring cell proton secretion (Klingler et al., 2002). The use of such a pH-sensitive biosensor allowed us to determine the effects of MDMA on the metabolic activation of myotubes. Transient exposition of myotubes cultured on micronets from three MHN individuals to 500 µM MDMA resulted in a phasic-tonic increase in the acidification rate. This effect was also almost abolished when the cells were exposed to -Bgt (Fig. 5).
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No Effects of MDMA and SCh on Isolated SR Vesicles. Caffeine and halothane are reagents known to release Ca2+ from the terminal cisternae of the SR, the most important myoplasmic Ca2+ store. We measured the effects of up to 170 µM ryanodine, 1200 µM halothane, 1000 µM SCh, and 700 µM MDMA on Ca2+ flux rates. In this preparation, which was taken from rat muscle, Ca2+ was released only upon incubation with halothane or ryanodine but not upon incubation with MDMA or SCh. This suggests that MDMA and SCh do not directly act upon RyR1 (Fig. 6).
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) (Fig. 7C).
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Chemical Analysis of Ecstasy Pills and MDMA Serum Levels. Of the 30 ecstasy pill samples confiscated by the police, free MDMA base content varied from 0.4 to 62.5%, with 25.5% being the mean MDMA base content. Three samples were found to be supplemented with caffeine. Of the 54 non-MDMA samples, the mean pure drug content of other amphetamine derivatives was 9.8%, and six were mixed with caffeine. MDMA levels of ecstasy users varied from 0.06 to 2.28 mg/l (0.3–11.7 µM) in 35 serum samples and were on average 0.40 mg/l (
2 µM). MDMA has a clearance of <12 h and a distribution volume of >4 l/kg. Peak concentration in muscle tissue may reach higher levels than in serum, especially if there is increased muscle perfusion (e.g., dancing). After intravenous administration of 1 to 2 mg/kg SCh during intubation, peak concentrations of SCh have been estimated to be 20 mg/l (100 µM). Because of the hydrophilic properties of SCh, muscular concentration may be somewhat lower (Hoshi et al., 1993).
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Discussion |
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; O'Connell and Heffron, 2000). In our analysis, about 10% of all pills (ecstasy and other amphetamines) were supplemented with caffeine, which acts as adenosine receptor antagonist and phosphodiesterase inhibitor. This may aggravate not only the stimulatory central but also the peripheral effects. This was obvious in our in vitro tests on normal adult skeletal muscle: although the highest concentration (1000 µM) was not able to induce a contracture by its own, the lowest [MDMA] used in our study, 100 µM, regularly increased its sensitivity for caffeine-induced contractures. This observation may be of clinical relevance because MDMA is often ingested in combination with fashion drinks ("energy drinks"), which contain up to 1 mg/ml caffeine. MDMA applied to cultured muscle cells induced intracellular Ca2+ peaks and acidification signals. The prevention of these effects by -Bgt, which binds with a high specificity and high affinity to the endplate, points to a direct interaction of MDMA with nAChR. The agonistic effect was verified by current measurements on HEK293 cells expressing 2 nAChR. Interestingly, a curare-like block by amphetamine overdosage was identified to cause paralysis (Skau and Gerald, 1978; Liu et al., 2003), pointing to the same target. In contrast to the effects of amphetamine, weakness has not yet been reported for illicit ecstasy users.
Both MDMA's inability to induce contractures on its own and its potentiating effect on caffeine- and halothane-induced contractures resemble the in vitro action of SCh (Galloway and Denborough, 1986; Ørding and Skovgaard, 1987; this study), one of the classical nAChR activators. Hence, we conclude that skeletal muscle is a target for MDMA outside the central nervous system that is affected via the nAChR of the neuromuscular junction. Mostly, strips taken from the quadriceps muscle do not contain an endplate. Nevertheless, MDMA can exert its activating effect on extrajunctional nAChR. Although the receptors of the adult type are less dense than at the endplate, they are frequent enough to produce measurable macroscopic currents (Koltgen and Franke, 1992). Also, primary myotube cultures express nAChR and their characteristics resemble the embryonic type (Lorenzon et al., 2002).
Are MHS Individuals Predisposed to Adverse MDMA Effects? Some of the clinical features such as metabolic acidosis, hyperkalemia, CK elevation, hyperthermia, and rhabdomyolysis in illicit MDMA users (Henry, 1992; Screaton et al., 1992; Hall et al., 1996) could be the result of a primary damage of skeletal muscle, e.g., MH (O'Leary et al., 2001). The response of such adverse MDMA effects to the MH antidote dantrolene supported this view (Hall et al., 1996), but the beneficial effects such as reduced myoplasmic Ca2+ and heat production are not restricted to MH crises (Hadad et al., 2005). We have tested the effects of MDMA on MHS muscle: None of the MHS muscles exhibited an in vitro contracture at 100 µM MDMA. This corresponds approximately to the highest MDMA serum value ever reported for an ecstasy user (De Letter et al., 2004), whereas the usual MDMA serum concentration ranged from 0.5 to 6 µM (Henry et al., 1992; this study). MDMA caused a smaller left-shift of the dose-response curve for caffeine- and halothane-induced contractures in MHS than in MHN muscle. Thus, MDMA increased the sensitivity of MHN muscle more than that of MHS. However, muscle bundles from three of eight MHS patients developed a contracture after exposure to 500 µM (n = 1), 1000 µM (n = 1), or 2000 µM (n = 1) MDMA. The smaller sensitivity increase for MHS than MHN muscle, the inconsistency, and the required high concentrations of MDMA-induced contractures do not support the view that muscle of MHS individuals is more sensitive to adverse MDMA reactions than normal muscle. Furthermore, we have excluded an MDMA-induced facilitation for the release of Ca2+ from the SR, the pathogenetic mechanism of MH (MacLennan and Phillips, 1992). To our knowledge there is no evidence that SCh triggers MH crises in humans in the absence of volatile anesthetics (Klingler et al., 2005).
In contrast to our results on excised human muscle, in vivo experiments on MHS pigs showed that intravenously injected MDMA (8–12 mg/kg b.wt. corresponding to a serum concentration of 7 µM) caused hyperthermia, acidosis and rhabdomyolysis (Fiege et al., 2003). Since MDMA leads to a serotonergic overstimulation, the MH-like symptoms in the pigs could be explained by a central serotonin syndrome (Gerbershagen et al., 2003).
Taking into account the central stimulation that activates the motoneuron system, an additional direct effect on nAChR may lead to relevant muscle symptoms even at low doses of MDMA. Since muscle fasciculations, muscle cramps, and muscle ache also have been typically reported by illicit MDMA users (Henry, 1992; Screaton et al., 1992; Hall et al., 1996), the underlying cause should be proximal to muscle, that means at the neuromuscular junction. nAChR are present in both the postsynaptic and at presynaptic parts, and from the latter, electric activity can spread out in retrograde direction along the motor unit and cause fasciculations and cramps (Guiloff and Modarres-Sadeghi, 1992). And all other muscle-related symptoms and signs of MDMA ingestion could be the result of postsynaptic nAChR stimulation, e.g., hyperkalemia, metabolic acidosis, muscle fiber swelling, and rhabdomyolysis could be simply explained by maldistribution of electrolytes due to K+ efflux and Na+ and Ca2+ influx through junctional and extrajunctional nAChR (Klingler et al., 2005).
Susceptibility to Adverse MDMA Reactions. MDMA may cause adverse effects in muscles abnormally reacting to nicotinic drugs. Particularly gross muscle with a high density of extrajunctional nAChR will be predisposed: generalized denervation processes such as in spinal muscular atrophies (Martyn et al., 1992; Lefebvre et al., 1998) or generalized muscle regeneration such as in progressive muscular dystrophies (Breucking et al., 2000; Gattenlohner et al., 2002). Ingestion of ecstasy also may be hazardous in patients with myotonic syndromes because the hyperexcitable muscle fiber membrane increases the effect of nicotinic agents on muscle (Lehmann-Horn and Jurkat-Rott, 1999).
In summary, MDMA 1) resembles the in vitro action of SCh on skeletal muscle strips, 2) induces Ca2+ transients in myotubes that are inhibited by -Bgt, 3) increases acidification and metabolism in myotubes, 4) has no effect on SR vesicles, and 5) activates nAChR dose dependently. Thus, the nAChR of the neuromuscular junction is one of the physiological targets of MDMA. This result was highly unexpected, although MDMA was reported to exert cholinergic effects in addition to serotonergic, dopaminergic, adrenergic, and histaminergic actions (Battaglia and De Souza, 1989; Crespi et al., 1997; Mechan et al., 2002; Gerbershagen et al., 2003). However, these cholinergic effects were ascribed to the muscarinic receptor types that are highly different from the nAChR at the neuromuscular junction. Our report draws attention to adverse ecstasy reactions particularly in patients with a generalized up-regulation of extrajunctional nAChR such as in some hereditary neuromuscular diseases. Retro- and prospective clinical observations will help reveal these potential pharmacogenetic reactions.
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Acknowledgements |
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Footnotes |
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Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.
ABBREVIATIONS: MDMA, 3,4-methylenedioxymethamphetamine; CK, creatine kinase; SCh, succinylcholine; MH, malignant hyperthermia; MHS, malignant hyperthermia-susceptible; IVCT, in vitro contracture test; GC, gas chromatograph; TFA, trifluoraceticanhydride; MHN, malignant hyperthermia-negative; RyR1, ryanodine receptor type 1; SR, sarcoplasmic reticulum; MOPS, 3-(N-morpholino)propanesulfonic acid; HEK, human embryonic kidney; nAChR, nicotinic acetylcholine receptor(s); -Bgt, -bungarotoxin; ACh, acetylcholine.
Address correspondence to: Dr. Frank Lehmann-Horn, Department of Applied Physiology, Ulm University, Albert-Einstein-Allee 11, 89069 Ulm, Germany. E-mail: frank.lehmann-horn{at}medizin.uni-ulm.de
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