Address correspondence and reprint requests to Frederic A. Meunier, Molecular Dynamics of Synaptic Function Laboratory, School of Biomedical Sciences, The University of Queensland, St Lucia, Brisbane, Queensland 4072, Australia. E-mail: firstname.lastname@example.org
Glycerotoxin (GLTx) is capable of stimulating neurotransmitter release at the frog neuromuscular junction by directly interacting with N-type Ca2+ (Cav2.2) channels. Here we have utilized GLTx as a tool to investigate the functionality of Cav2.2 channels in various mammalian neuronal preparations. We first adapted a fluorescent-based high-throughput assay to monitor glutamate release from rat cortical synaptosomes. GLTx potently stimulates glutamate secretion and Ca2+ influx in synaptosomes with an EC50 of 50 pm. Both these effects were prevented using selective Cav2.2 channel blockers suggesting the functional involvement of Cav2.2 channels in mediating glutamate release in this system. We further show that both Cav2.1 (P/Q-type) and Cav2.2 channels contribute equally to depolarization-induced glutamate release. We then investigated the functionality of Cav2.2 channels at the neonatal rat neuromuscular junction. GLTx enhances both spontaneous and evoked neurotransmitter release causing a significant increase in the frequency of postsynaptic action potentials. These effects were blocked by specific Cav2.2 channel blockers demonstrating that either GLTx or its derivatives could be used to selectively enhance the neurotransmitter release from Cav2.2-expressing mammalian neurons.
The high specificity of the action of neurotoxins has been a great asset in the deciphering of neurophysiological mechanisms at the molecular level. Some neurotoxins have found their place in the pharmacopoeia, such as curare derivatives, while a number are awaiting the outcome of clinical trials (Foran et al. 2003). The vast majority of these neurotoxins act by blocking the activity of their targets. A few neurotoxins such as α-latrotoxin have shown stimulatory properties, and have given new functional insights on physiological processes such as exocytosis and endocytosis (Schiavo et al. 2000). However, these neurotoxins commonly exhibit irreversible paralytic actions resulting from the depletion of presynaptic vesicle pools (Ceccarelli and Hurlbut 1980; Tsang et al. 2000; Rigoni et al. 2004), therefore limiting their therapeutic potential.
We have recently identified and purified a novel excitatory neurotoxin from Glycera convoluta venom; the 320-kDa neurotoxin named glycerotoxin (GLTx) (Meunier et al. 2002). GLTx reversibly increases both spontaneous and evoked quantal acetylcholine (ACh) release at the frog neuromuscular junction (NMJ) by acting selectively on N-type Ca2+ (Cav2.2) channels (Meunier et al. 2002). Whole-cell patch-clamp analysis revealed that GLTx promotes a leftward shift in the current–voltage relationship of the channels, thereby making this neurotoxin the first Cav2.2 channel activator that has been described (Meunier et al. 2002).
In this study, we demonstrate that GLTx stimulates neurotransmitter release from developing motor nerve terminals known to transiently express Cav2.2. In order to test the potential of GLTx in stimulating glutamate release from central neurons, we have adapted a fluorescence-based glutamate release assay from rat brain synaptosomes to a high-throughput 96-well plate format. Using this assay, we found that GLTx at picomolar concentrations stimulates glutamate release from rat cortical synaptosomes via Cav2.2 channels. Either the dysfunction or the impairment of the activities of voltage-gated Ca2+ channels have been linked to several medical conditions (Fletcher et al. 1996; Ophoff et al. 1996; Kraus et al. 2000; Wappl et al. 2002). By specifically interacting with Cav2.2 channels in the mammalian CNS, GLTx represents a novel tool for increasing neurotransmitter release from Cav2.2 channel-expressing neurons.
Materials and methods
Reagents and toxins
ω-CTx-GVIA and ω-CTx-MVIIA were purchased from Alomone (Jerusalem, Israel), ω-CTx-CVID was kindly donated by R. Lewis (Institute for Molecular Biosciences, Brisbane, Australia), fluo-3 and fura-2/AM were obtained from Molecular Probes (Invitrogen, Mount Waverly, Australia). GLTx was prepared as previously described and tested for its ability to stimulate catecholamine release from bovine adrenal chromaffin cells (Meunier et al. 2002). All other reagents were from Sigma (Castle Hill, NSW, Australia).
Left and right hemidiaphragm muscles with their associated phrenic nerves were isolated from neonatal rats (day 3–4) killed by dislocation of the cervical vertebrae. Animals were treated in accordance with animal ethics guidelines of the University of Queensland. The two hemidiaphragms were separated and each was mounted in a Rhodorsil®-lined organ bath (Rhône-Poulenc, St Fons, France) (2 mL) superfused with a physiological solution of the following composition (mm): NaCl 154; KCl 5; CaCl2 2; MgCl2 1; HEPES buffer 5; glucose 11; pH 7.4; gassed with pure O2. Miniature end-plate potentials (MEPPs) were recorded from end-plate regions at room temperature (22–24°C), with intracellular microelectrodes filled with 3 m KCl (of 20–50 MΩ resistance), using conventional techniques and an Axoclamp-2A system (Molecular Devices, Surrey Hills, Victoria, Australia). Recordings were made from different end-plates at indicated times prior to and throughout the application of GLTx. Amplified electrical signals were collected and digitized at a sampling rate of 25 kHz, with the aid of a computer equipped with an analogue–digital interface board. Data were collected and analysed with Scan software, kindly provided by Dr John Dempster (University of Strathclyde, Glasgow, UK), running on a PC equipped with an analogue–digital converter (Model DT2821; Data Translation, Waltham, MA, USA).
Extracellular recordings were made from end-plate regions at room temperature (22–24°C). Extracellular electrodes (tip diameters 10–15 µm), filled with Tyrode's solution (in mm: NaCl 136.7; KCl 2.68; NaH2PO4 1.75; NaHCO3 16.3; MgCl2 1.0; d-glucose 7.8; CaCl2 2; pH 7.4), were used to record nerve terminal impulses, evoked excitatory postsynaptic currents (EPC) and spontaneous excitatory postsynaptic currents (SEPC). Recordings were made in the presence of tetrodotoxin (TTX; 1 µm) from different end-plates at indicated times prior to and throughout the application of GLTx. The phrenic nerve was drawn into a glass pipette that was filled with Tyrodes solution and gassed with carbogen (95% O2/5% CO2). The nerve was stimulated using two silver chloride electrodes. Square-wave pulses with amplitude of 20 V and duration 0.08 ms were used to stimulate the nerve at a frequency of 0.2 Hz. The resulting amplified electrical signals were collected and digitized at a sampling rate of 10 kHz, with the aid of a computer equipped with an analogue–digital interface board (MacLab4, Natick, MD, USA). Data were collected and analysed using Scope and Chart software (MacLab4). The quantal content was estimated by calculating the EPC/SEPC ratio. The extracellular electrode was carefully positioned over the end-plate region and repositioned until the SEPCs were maximal in amplitude; this was close to the point were the axon makes contact with the skeletal muscle fiber. The frequency of SEPCs was monitored throughout this procedure to ensure that the positioning of the electrode over the end-plate did not damage the nerve terminal, and that the pressure did not alter the SEPC frequency nor evoke transmitter release.
Synaptosomes were prepared as described (Dunkley et al. 1988; McMahon et al. 1989) with a few adaptations. Briefly, eight-week-old male Wistar rats were killed by cervical dislocation and their brains were removed and placed immediately into ice-cold sucrose buffer (0.32 m sucrose, 20 mm HEPES-NaOH, pH 7.3). The brainstem, cerebellum, meninges and white matter were removed and the intact cortex was homogenized in 9 mL of ice-cold sucrose buffer plus protease inhibitors (Roche Diagnostics, Castle Hill, NSW, Australia) using a Teflon-glass potter homogenizer (12 strokes, 400 rpm). The homogenate was centrifuged at 1000 g for 10 min at 4°C. The supernatant was collected and diluted to 14 mL and separated on discontinuous percoll gradients (3, 10, 15 and 23%) by centrifugation at 32 500 g for 6 min at 4°C. Layer 4 at the interface of the 15 and 23% Percoll layers was collected and diluted 10 times with HEPES-buffered media (HBM; mm): NaCl 140; KCl 5; HEPES-NaOH 20; NaHCO3 1.5; MgCl2 10; Na2HPO4 1.2, d-glucose 10; pH 7.4. Synaptosomes were washed twice by centrifugation at 16 000 g for 10 min at 4°C, resuspended in 4 mL of HBM and kept on ice until required. The protein concentration was determined using a Bradford assay (Bio-Rad Laboratories, Hercules, CA, USA).
Glutamate release assay
Glutamate release was determined fluorometrically based on a method previously described (McMahon et al. 1989) and adapted to a 96-well plate format (Sim et al. 2005). Briefly, 50 µg of synaptosomes were pre-incubated with 2.5 mm NADP+ and 13.5 U of glutamate dehydrogenase in HBM. NADPH fluorescence produced by the conversion of glutamate to 2-oxoglutarate was monitored at 1-s intervals using the FLUOstar OPTIMA 96-well microplate reader equipped with two independent injecting devices operated by star Fluostar Optima software (v1.32; BMG Labtechnologies, Mornington, Victoria, Australia). Depolarizing agents were applied via injectors as indicated. When appropriated, synaptosomes were pre-treated with ω-CTx-GVIA (1 µm) for 10 min prior to stimulation (either with GLTx or with KCl). Glutamate release was expressed as ΔF/F0, where F0 represents the initial baseline fluorescence, and was converted in nmol glutamate released/mg of protein.
Ca2+ measurements using a microplate reader
Synaptosomes were incubated with fura-2 plus pluronic acid (1 µm) on ice for 30 min, washed once with HBM and then resuspended at 1 mg/mL in HBM containing Ca2+ (2 mm). Fluorescence recordings were taken at 1.43-s intervals by alternating the excitation wavelengths of 340 and 380 nm and monitoring the emission wavelength at 510 nm using the FLUOstar OPTIMA microplate reader. The fluorescence ratio (340/380 nm) was expressed as ΔF/F0 versus time in seconds.
Motor neuron preparation and Ca2+ imaging
Spinal cord motor neurons were purified from mice embryos (at day 13) and plated on glass coverslips coated with a mixture of polyornithine and collagen as previously described (Hafezparast et al. 2003). Cells were loaded with the acetoxymethyl ester form of either fura-2 or fluo-3 [5 µm with 0.02% pluronic acid (w/v) for 45 min at 37°C in the dark] in HBM. After washing and incubation at 37°C for 15 min, the dishes were placed in an environmental chamber (37°C) on the stage of either a confocal microscope (LSM510; Zeiss, Standort Göttingen, Germany) equipped with an Acroplan 60× (NA 0.9) water-immersion lens or an epifluorescent microscope (Diaphot 300; Nikon, Kingston upon Thames, UK) equipped with either a Plan Fluor 60× or a 100× (NA 1.3) oil-immersion objective. Fura-2-loaded cells were excited through 340- and 380-nm filters and images were acquired using a cooled CCD camera (Orca I; Hamamatsu City, Shizuoka Pref, Japan) controlled by Aqm2000 software (Kinetic Imaging, Wirral, Merseyside, UK). Images from either fura-2 or fluo-3-loaded neurons were taken at 4-s intervals and ratios were obtained using either Lucida (v.4.0; Kinetic Imaging) or Lsm510 software, respectively. F/F0 was calculated as previously described (Meunier et al. 2002).
GLTx binding assay
The crude synaptic plasma membrane fraction (LP1; 100 µg; Huttner et al. 1983), was diluted in ice-cold binding buffer (mm: NaCl 62.5; HEPES-NaOH 10; EDTA 0.1; EGTA 1.0; bovine serum albumin, BSA 0.1%; pH 7.2) plus protease inhibitors and incubated with increasing concentrations of GLTx, heat-treated GLTx (3 min, 100°C) or ω-CTx-MVIIA (Alomone) as indicated for 10 min prior to the addition of 0.1 nm radiolabelled ω-CTx-GVIA (NEN, Hounslow, UK). The mixture was left for 1 h at 4°C under gentle agitation, transferred onto a 25-mm diameter GF/C filter (1822025; Whatman, Maidstone, Kent, UK) pre-soaked with ice-cold binding buffer containing 0.5% (v/v) polyethyleneimine, washed twice with an excess of ice-cold binding buffer and counted (Wallac 1450 MicroBeta; Perkin-Elmer, Fremont, CA, USA). Samples containing equal quantities of protein (50 µg) were analysed for the presence of Cav2.2 by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and western blotting using anti-Cav2.2 (Chemicon, Boronia, Victoria, Australia).
Statistical analysis was performed using the Student's t-test (two-tailed, unpaired). Values are expressed as mean ± SEM and data were considered significant at *p < 0.05 and at **p < 0.01.
To ascertain whether GLTx was active on mammalian neurons, we developed a high-throughput version of a previously described assay (McMahon et al. 1989) in order to detect endogenous glutamate release from rat-brain synaptosomes using a 96-well plate reader (Sim et al. 2005). One of the main advantages of this technique is the possibility to make concentration–dependence studies, which were unattainable using earlier methodology (McMahon et al. 1989). We first tested the validity of our assay by measuring the release of endogenous glutamate from synaptosomes triggered by depolarization (Fig. 1a). We then used 4-aminopyridine (4-AP), a potassium channel blocker, in order to prevent synaptosome repolarization and promote an increase in the presynaptic Ca2+ channel opening time (Van der Kloot 1994). 4-AP promotes glutamate release and potentiates the depolarization-evoked response (Fig. 1a). Picomolar concentrations of GLTx elicited glutamate release comparable to that evoked by depolarization (Fig. 1b). This effect was strictly dependent on the presence of external Ca2+ (Fig. 1b).
The concentration response indicated that GLTx stimulates glutamate release with an EC50 of 50 pm (Figs 1c and d). These experiments establish that GLTx is a potent secretagogue, capable of stimulating glutamate release from purified synaptosomes at picomolar concentrations.
To examine the involvement of Cav2.2 channels in mediating the stimulatory effect of GLTx on mammalian brain synapses, we analysed the contribution of different Cav2 channels in depolarization-induced glutamate release. Previous methodologies have generated disparate results as to the relative contributions of Cav2.2 and Cav2.1 channels in regulated secretion (Turner et al. 1993; Vazquez and Sanchez-Prieto 1997; Leenders et al. 1999, 2002). We tested the sensitivity of synaptosomes to the novel selective Cav2.2 channel inhibitor ω-CTx-CVID (Adams et al. 2003) by pre-incubation with ω-CTx-CVID (1.5 µm) followed by depolarization. A 45 ± 5% inhibition of depolarization-induced glutamate release was detected, indicating that Cav2.2 channels have a major role in glutamate release from cortical synaptosomes (Fig. 2a). A similar inhibition was observed with the Cav2.2 channel blocker ω-CTx-GVIA (data not shown). The contribution of Cav2.1 channels in depolarization-induced glutamate release was also tested using agatoxin-IVA (2 µm) (Turner et al. 1993), which inhibited glutamate release by 52 ± 6%. Glutamate release was further inhibited by the combined treatment with both ω-CTx-CVID and agatoxin-IVA (Fig. 2a), indicating that both Cav2.2 and Cav2.1 channels contribute to this process.
Next, we examined whether GLTx was acting exclusively on Cav2.2 channels as previously described in other preparations (Meunier et al. 2002). Synaptosomes were pre-incubated with ω-CTx-GVIA (1 µm) and stimulated with GLTx (65 pm). Under these conditions, the stimulatory effect of GLTx was abolished (Fig. 2b). Importantly, agatoxin-IVA pre-incubation (2 µm) did not affect GLTx-induced glutamate release (Fig. 2b).
We then investigated whether ω-CTx-GVIA-mediated inhibition of the stimulatory effect of GLTx could result from a competition between these two neurotoxins for a common binding site on the Cav2.2 channel. We first checked the presence of Cav2.2 channels in rat brain subcellular fractions by western blotting using an anti-α1B antibody. The α1B subunit was detected in both a synaptosomal extract and a synaptic plasma membrane fraction (LP1) (Fig. 3a). LP1 fractions were then incubated with increasing concentrations of either native or heat-inactivated GLTx for 10 min before the addition of [I125]ω-CTx-GVIA. Following 1 h of incubation, membranes were immobilized on Whatman filters, washed and processed for radioactive counting. Neither native nor heat-inactivated GLTx was capable of inhibiting [I125]ω-CTx-GVIA binding (Fig. 3b). A control experiment was performed using ω-CTx-MVIIA, which is known to compete with ω-CTx-GVIA for the same binding site. As expected, increasing concentrations of ω-CTx-MVIIA abolished ω-CTx-GVIA binding to Cav2.2 channels as previously established (Fig. 3c) (Kristipati et al. 1994). These data demonstrated that at physiological concentrations, GLTx and ω-CTx-GVIA interact with the Cav2.2 channel on distinct binding sites.
To test whether the secretagogue activity of GLTx results from Ca2+ influx in synaptosomes, fura-2-loaded synaptosomes were treated with GLTx (Fig. 4a). We found that GLTx (65 pm) was indeed capable of inducing a transient increase in intracellular Ca2+, which can be blocked by ω-CTx-GVIA (Figs 4a–c). The GLTx-induced Ca2+ increase was concentration dependent with an apparent EC50 of 57 pm (Fig. 4d), which is in good agreement with that seen for GLTx-induced glutamate release (Fig. 1d).
Combined with our previous results, these data suggested that GLTx promotes a strong secretagogue activity from Cav2.2-expressing neurons.
Mature rat NMJs are known to mainly express Cav2.1 channels (Protti and Uchitel 1993; Katz et al. 1997; Rahamimoff et al. 1999) and are therefore insensitive to GLTx (Meunier et al. 2002). In contrast, neonatal motor nerve terminals were shown to transiently express functional Cav2.2 channels (Rosato-Siri and Uchitel 1999; Rosato-Siri et al. 2002) and should therefore be sensitive to GLTx. As shown in Fig. 5(a), this is indeed the case. Upon the addition of GLTx (150 pm), a marked increase in the frequency of MEPP was observed as compared with control (Figs 5a and b). A GLTx-induced increase in MEPPs frequency was observed for at least 1 h and was reversible by washing (Fig. 5a). There was no noticeable change in the monoquantic MEPP amplitude, rise time or exponential decay (data not shown). We next tested whether the stimulatory activity of GLTx could be abolished by ω-CTx-MVIIA. As shown in Fig. 5(b), ω-CTx-MVIIA not only reversed the stimulatory effect of GLTx, but also prevented it, demonstrating that GLTx acts via Cav2.2 channels at the neonatal rat NMJ.
In the absence of TTX, spontaneous twitching was observed upon the addition of GLTx at the neonatal NMJ, suggesting that the level of neurotransmitter release elicited by GLTx was high enough to promote postsynaptic action potentials. To investigate the effect of GLTx on evoked neurotransmitter release, we used extracellular recording to monitor EPC amplitudes upon incubation of the neonatal NMJ with 150 pm GLTx. This treatment promoted an increase in EPC amplitudes (Fig. 6a) with no significant change in SEPC (data not shown). Consequently, GLTx increased the mean quantal content by about two-fold (Fig. 6b). This effect on phasic release was completely reversible by washing (Fig. 6b). Moreover, the rise in quantal content observed with GLTx was large enough to double the appearance of postsynaptic action potentials (Figs 6c and d).
To confirm that the increase in spontaneous release upon GLTx treatment was caused by an increase in intracellular Ca2+, we tested the effect of GLTx on purified motor neurons isolated from rat spinal cord. Motor neurons have been shown to express functional Cav2.2 channels (Scamps et al. 1998). Neurons were loaded with fluo-3 and examined by confocal microscopy. GLTx application (150 pm) promoted spontaneous oscillations in intracellular free Ca2+ followed by an overall increase in the intracellular Ca2+ signal (Figs 7a and b). This experiment was repeated using fura-2-loaded neurons with identical results (data not shown). Importantly, the inhibition of Cav2.2 by pre-incubation of motor neurons with ω-CTx-CVID completely prevented the increase in intracellular Ca2+ (Figs 7a and b).
Altogether, these experiments demonstrate that GLTx promotes neurotransmitter release by acting on N-type Ca2+ channels and allowing Ca2+ influx in motor nerve terminals.
GLTx is one of the few known toxins capable of stimulating secretion directly and reversibly, allowing unique functional insights into the study of exocytosis. In this work, we demonstrated that GLTx stimulates neurotransmitter release from neonatal NMJs and adult CNS synapses via Cav2.2 channels.
GLTx differs substantially from these neurotoxins because it stimulates exocytosis by selectively up-regulating Cav2.2 channels. A direct interaction between GLTx and Cav2.2 has been demonstrated previously by immunoprecipitation (Meunier et al. 2002). In this study, we aimed to find whether GLTx and ω-CTx-GVIA share the same binding site on Cav2.2 channels. The binding site for ω-CTx-GVIA has been well-characterized and lies on the extracellular loop between the IIIS5 and IIIH5 transmembrane domains, suggestive of a pore-blocking mechanism (Ellinor et al. 1994; Feng et al. 2001, 2003). At concentrations relevant for its action, GLTx is unable to displace bound ω-CTx-GVIA, suggesting that the two neurotoxins bind to different sites on Cav2.2 channels.
The contribution of several members of the Cav2 family in mediating neurotransmitter release in the brain has been extensively investigated. Although glutamate secretion was originally thought to rely uniquely on Cav2.1 channels (Turner et al. 1993), more recent studies have challenged this view (Vazquez and Sanchez-Prieto 1997; Leenders et al. 1999). Despite the fact that all these studies have analysed glutamate release using synaptosomes preloaded with radioactive glutamate, subtle variations between the uptake protocols may explain the different results. The development of a high throughput synaptosome assay measuring the release of endogenous glutamate allowed us to bypass these experimental problems and investigate the contribution of Cav2.1 and Cav2.2 channels in mediating depolarization-evoked glutamate secretion. As shown in Fig. 2, we found a comparable contribution of both channels to this process, suggesting that both Cav2.1 and Cav2.2 are functionally coupled to the synaptic release machinery in rat brain synaptosomes.
GLTx increases both spontaneous and evoked quantal release via the Cav2.2 channel
The increase in quantal content observed at the rat neonatal NMJ and the frog NMJ is likely to reflect a rise in the intraterminal Ca2+ level. GLTx has been shown to directly interact with Cav2.2 channels and to promote Ca2+ influx in neurosecretory cells (Meunier et al. 2002), cortical synaptosomes and purified motor neurons (this work). The sustained increase in spontaneous release triggered by GLTx is likely to stem from poorly characterized oscillations in Ca2+ levels also observed in GLTx-treated neurons and chromaffin cells (data not shown). It is worth noting that the increase in quantal content observed at the neonatal rat NMJ is less pronounced than that observed at the frog NMJ (Meunier et al. 2002). Indeed, GLTx increases the amplitude of the evoked response at the adult frog NMJ by up to 20-fold, whereas only a two-fold increase was detected at the neonatal NMJ. This might reflect a smaller number of functional presynaptic Cav2.2 channels only transiently expressed after birth. Alternatively, such a difference could also result from an incomplete coupling of the Cav2.2 channel with the release machinery. A poor degree of co-localization between Cav2.2 channels and release sites could account for the difference in Ca2+ sensitivity observed between the two NMJ preparations. A reduced paired-pulse facilitation has recently been described in the Cav2.1 knockout known to express Cav2.2 as a compensatory mechanism (Pagani et al. 2004).
Altogether, these findings confirm that GLTx is not only capable of stimulating exocytosis from catecholamine-containing large dense core vesicles (Meunier et al. 2002) but also from synaptic vesicles containing either glutamate or ACh in different synapses.
The mode of action of GLTx
GLTx is a very active neurotoxin that stimulates neurosecretion at picomolar doses via a selective interaction with Cav2.2 channels. This binding promotes a leftward shift in the current–voltage relationship of Cav2.2 (Meunier et al. 2002), which could ultimately be responsible for GLTx activity. Surprisingly, the observed rate of onset of GLTx action is much faster than that expected from three-dimensional free diffusion, given the low concentration at which GLTx is active (50–100 pm) and its large molecular weight (320 kDa). This argues against a classic ligand-effector equilibrium for the GLTx-Cav2.2 binding and suggests a possible indirect action of GLTx on the channel. Several hypotheses may explain this discrepancy. An attractive possibility is that GLTx possesses (or stimulates) a still uncharacterized enzymatic activity generating a fast diffusible metabolite responsible for Cav2.2 channel opening. In this model, a membrane-directed activity, such as a phospholipase, could potentially modify the lipid micro-environment of the channel with consequences on its biophysical properties. However, studies on snake neurotoxins bearing a phospholipase A2 activity have shown relatively slow initial kinetics and poor reversibility (Montecucco et al. 2004; Rigoni et al. 2004), two characteristics that do not match the observed features of GLTx. Moreover, early studies on the partial purification of G. convoluta have shown that the stimulatory effect on transmitter release is found in a high-molecular-weight fraction distinct from those containing protease and phospholipase activities (Thieffry et al. 1982). Finally, pronase treatment of synaptosomes prevented the effect of GLTx, reinforcing the idea that this toxin binds to an externally accessible protein of the plasma membrane (Morel et al. 1983).
Alternatively, the onset of GLTx action on Cav2.2 channels might be hastened by its partition at the surface of the plasma membrane, thereby shifting the diffusion problem from three to two dimensions. In this regard, GLTx is a very large toxin, expected to have many charged amino-acid residues that may influence the local charge of the lipid bilayer. This could help in concentrating GLTx near the Cav2.2 channel and facilitate their interaction. Further knowledge on the sequence of GLTx and its surface charge distribution will be crucial to gain insights on the mode of action of this novel neurotoxin.
CNS localization of Cav2.2 channels and potential therapeutic use of GLTx
The localization of Cav2.2 channels in the adult rat brain has been previously characterized. In the cerebral cortex and hippocampus, Cav2.2 channels are mainly localized along dendrites, to a lesser extent on their cell bodies and in clusters at presynaptic nerve terminals (Westenbroek et al. 1992). In the forebrain and midbrain, Cav2.2 channels mainly associate with dendrites and in some cases cell bodies, whereas in the cerebellum they are primarily found on the proximal part of the dendritic tree of Purkinje cells (Westenbroek et al. 1992).
Either the dysfunction or impairment of calcium channels have been linked to several medical conditions. For example, mutations in the Cav2.1 genes have been reported to be involved in absence seizures, cerebellar ataxia and migraine (Fletcher et al. 1996; Ophoff et al. 1996; Kraus et al. 2000; Wappl et al. 2002). These mutations result in a reduction of current density in cerebellar Purkinje cells and CA3 pyramidal cells in the hippocampus (Pietrobon 2002). As Cav2.2 channels are functionally expressed in these cells, GLTx may be able to increase Ca2+ currents and therefore compensate for the loss of Cav2.1 channels. This suggests a potential use for GLTx in therapeutic applications aiming at increasing neurotransmitter release from Cav2.2-expressing neurons.
We are indebted to Dr S. Osborne for her critical reading of the manuscript and the preparation of LP1, and to Dr G. Lalli for the preparation of spinal cord neurons. This work was supported by ARC (DP0452106 to FAM and LR), by Cancer Research UK (GS), by an operating grant from the Heart and Stroke Foundation of Alberta and the North-west Territories (to GWZ). GWZ is a CIHR Investigator and a Senior Scholar of the Alberta Heritage Foundation for Medical Research. DP was supported by The University of Queensland Post-Graduate Research Scholarship.