Calcium currents from jellyfish striated muscle cells: preservation of phenotype, characterisation of currents and channel localisation

Calcium ions play an important role in regulating a number of cellular processes, with the calcium ion concentration in various compartments determining the onset, speed and termination of various biochemical and biophysical events (Berridge, 1993). Voltage-gated calcium channels play a crucial role since their activity will influence the cytoplasmic concentration of calcium ions. Among the numerous examples of the regulatory activity of Vg calcium channels are smooth and striated muscle contraction, transmitter release at synapses, signal transduction, development of vascular tone, release of hormones and cellular growth (Hille, 1992).

Voltage-gated calcium channels can be categorised into high-voltage-activated (HVA) and low-voltage-activated (LVA) channels. In general, HVA calcium channels activate above –40 mV. The activation and inactivation kinetics of HVA calcium channels are quite variable but they can be identified by their sensitivity to various pharmacological agents. They are categorised into at least five classes, namely L, N, P, Q and R types, based on their pharmacological properties. LVA channels activate at around –70 to –40 mV, inactivate rapidly, and are sensitive to nickel and mibefradil (Bean, 1985; Akaike et al., 1989; Huguenard, 1996; Ertel et al., 1997). To date, only one type of LVA channel has been identified, the T-type calcium channel.

Voltage-gated calcium channels are found in all muscle types; however, their roles in muscle contraction differ depending on the muscle type. Influx of external calcium ions is required to initiate contraction in both smooth and cardiac muscle but not in skeletal striated muscle of vertebrates (Bers, 1991), where depolarisation of the membrane triggers release of calcium from the sarcoplasmic reticulum (Endo, 1977). All cardiac myocytes possess L-type calcium channels, while T-type channels are also present in cardiac cells that display primary or secondary pacemaker activity such as at the sinoatrial node, sinus venosus and Purkinje fibres (Hagiwara et al., 1988; Hirano et al., 1989; Tseng and Boyden, 1989; Bois and Lenfant, 1991). However, T-type channels are absent or present at low densities in ventricular myocytes (Bers, 1991; Campbell and Strauss, 1995). Thus, the primary role of L-type calcium channels appears to be the control and modulation of the electromechanical coupling process, while T-type channels are involved in pacemaker activity.

Several voltage-gated calcium currents have been recorded from neurones in jellyfish (Mackie and Meech, 1985; Anderson, 1987; Przysiezniak and Spencer, 1992), smooth muscles in ctenophores (Bilbaut et al., 1988; Dubas et al., 1988), and fertilised eggs in ctenophores (Barish, 1984). Calcium currents are present in the striated swimming muscle of the medusa Polyorchis penicillatus (Spencer and Satterlie, 1981). More recently, using isolated striated muscle strips from this jellyfish, we have shown that contraction depends on extracellular calcium and can be inhibited by the calcium channel blocker nitrendipine (Lin et al., 2000). Using both electrophysiological techniques and specific labelling, we report the presence of low-voltage-activated calcium currents in striated myocytes of the hydrozoan jellyfish Polyorchis penicillatus. Retention of functional channels in these cells required dissociation at 30°C, with much of the current being lost at room temperature. A primary function of calcium currents through this channel population is to initiate muscle contraction. We have determined that the muscle feet have a far higher density of these channels than the somata.

Medusae of Polyorchis penicillatus were collected from Bamfield Inlet or Pachena Bay, near Bamfield, BC, Canada, and maintained in an aquarium with running sea water (9–12°C) at the Bamfield Marine Station, Bamfield, BC, Canada. The normal water temperature that these jellyfish experienced ranged from 7 to 20 °C throughout the year.

Subumbrellar sections were excised from the whole jellyfish and the radial and ring canals trimmed away. The muscle layer was stripped off the endodermal lamella using a pair of forceps. One medium-sized jellyfish (about 15–20 mm in bell diameter) yielded approx. 200 μl of striated muscle cells with some mesoglea attached. The stripped tissues were put into a 1.5 ml Eppendorf tube and a digestion solution of 1 mg ml^–1 Pronase (Boehringer Mannheim) dissolved in artificial sea water (ASW), either prewarmed to 30°C or at room temperature (20–22°C), was added to the Eppendorf tube to a final volume of 1 ml. Test tubes were left undisturbed at either room temperature (20–22°C) or at 30°C for 15–20 min. After digestion, a few gentle taps on the test tube was all that was needed to separate the tissue into single muscle cells. Muscle cells were dissociated at 30°C because the contractile phenotype appeared to be fixed by heat. Muscle cells dissociated at 30°C were easily identified by their long contractile processes, whereas muscle cells dissociated at room temperature could be identified by the lack of muscle feet and the retracted myofibres within the cytoplasm. Any remaining endodermal cells could be identified by their cuboidal or columnar shape. Before recording, cells were washed with the appropriate bath solution three times and resuspended in bath solution to a final volume of 500 μl.

Recordings were made immediately after dissociation. Each batch of cell suspension was used for about 3–4 h. One or two drops of cell suspension were added into a stagnant perfusion chamber and then cells were allowed to settle (some remained suspended) before recording.

Electrophysiological recording and data analysis followed the technique of Grigoriev et al. (Grigoriev et al., 1996). Briefly, whole-cell, tight-seal recordings were made using borosilicate glass pipettes (TW-150-4, World Precision Instruments), which were made on a Sutter automated puller, with resistances of 2–3 MΩ when filled with the pipette solution. Recordings were made with an Axopatch-1D amplifier (Axon Instruments), low-pass filtered at 3 kHz using a 4-pole Bessel filter, and digitised using a Labmaster TL-125 acquisition board (Axon Instruments). Individual cells were viewed under phase contrast with a Nikon Diaphot inverted microscope. A piezo-electric driver (Burleigh) was used to manipulate the pipette onto the cell surface. Perfusion with the relevant external recording solution started after successful seal formation. No contraction was observed during recording. Stimulus control, data acquisition and analyses were performed with pCLAMP 6.0 software (Axon Instruments) on a Dell personal computer. Leakage and capacitative currents were subtracted, prior to test pulses, using –P/4 or –P/5 protocols from a holding potential of –80 mV (pCLAMP 6.0). Series resistance (R_s) was compensated optimally to minimise voltage errors, and was usually set to values of ≥80 %. C_m (membrane capacitance) and R_s were obtained by minimising the capacitative transient in response to a hyperpolarising voltage step. Mean R_s was 4.39±0.81 ΜΩ (N=20) before compensation, and mean C_m was 5.49±1.07 pF (N=20) before compensation. All experiments were carried out at room temperature (20–22°C).

Before use all recording solutions were filtered through cellulose acetate membrane cartridges with a 0.2 μm pore size. ASW at pH 7.5 contained: NaCl 376 mmol l^–1, Na₂(SO₄) 26 mmol l^–1, MgCl₂ 41.4 mmol l^–1, CaCl₂ 10 mmol l^–1, KCl 8.5 mmol l^–1 and N-2-hydroxy-ethylpiperazine-N′-2-ethanesulphonic acid (Hepes) hemisodium salt 10 mmol l^–1. The bath solution at pH 7.5 contained: MgCl₂ 40 mmol l^–1, CaCl₂ (or BaCl₂ or SrCl₂) 10 mmol l^–1, KCl 15 mmol l^–1, N-methyl glucamine-HCl (NMG-Cl) 429 mmol l^–1, Hepes (free acid) 10 mmol l^–1. The pipette solution (pH 7.5) used for recording total membrane currents contained: NaCl 50 mmol l^–1, MgCl₂ 2 mmol l^–1, CaCl₂ 1 mmol l^–1, ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid (EGTA) 11 mmol l^–1, KCl 350 mmol l^–1, KOH 30 mmol l^–1, Hepes (free acid) 10 mmol l^–1. The pipette solution (pH 7.5) used for isolating calcium currents contained: NaCl 50 mmol l^–1, MgCl₂ 2 mmol l^–1, CaCl₂ 1 mmol l^–1, EGTA 11 mmol l^–1, CsCl 350 mmol l^–1, CsOH 30 mmol l^–1, Hepes (free acid) 10 mmol l^–1, TEA-Cl 20 mmol l^–1. Using the latter pipette solution all outward currents were blocked totally within the first minute after break-through.

The considerable thickness and elasticity of the mesoglea in the bell region made muscle tension recordings from this region extremely variable. A preliminary study indicated that muscle strips from bell regions and vela (the shelf of tissue at the opening of the bell) had similar pharmacologies. Therefore we used muscle strips from vela, which had a thin lamina of mesoglea, to measure tension development. The vela of medusae, anaesthetised in 1:1 isotonic MgCl₂ (0.33 mol l^–1) and ASW, were excised so as to provide continuous strips of maximal width. Care was taken not to include portions of the innervating motor-neurone network in the strips by bisecting the strip lengthwise and using only the half closest to the free margin. The free ends of each velar strip were pinned to the Sylgard base of a 35 mm Petri dish, which also contained a pair of embedded Ag/AgCl₂ stimulating electrodes connected to a Grass S44 stimulator. The velar strip ran between the two stimulating electrodes and around a small hook attached to a capacitative force transducer (Kent Scientific Corporation). Tension on the strip was adjusted using a micromanipulator so as to remove any slack in the muscle strip. Repetitive square pulses of 30 ms duration and 0.2 Hz were given to produce a train of twitch contractions that did not show fatigue. The stimulation voltage was determined by increasing the voltage until no increase in the amplitude of contraction at the stimulation frequency of 0.1 Hz occurred. The voltages used usually were between 30 and 40 V. Perifusion was by a peristaltic pump at 1.5 ml min^–1 and the perifusate was removed by vacuum. All perifusates were kept at 12–14°C during the experiment by running part of the perfusion tubing through an ice bucket. Transduced tension was recorded on a digital, Dash-IV pen-recorder (Astro-Med Inc.). The amplitude of contractile force for each condition (control, drug effect and wash) was calculated by averaging 30 individual twitch contractions.

The method used to localise calcium channels using dihydropyridine-BODIPY (fDHP) followed the protocol of Schild et al. (Schild et al., 1995), with some modification. Dissociated cells were incubated with both fDHP and the styryl dye, RH414 (Molecular Probes, Inc.) at final concentrations of 10 μmol l^–1 and 5 μmol l^–1, respectively, for 15 min at room temperature before examination. A laser-scanning, confocal microscope (Molecular Dynamic, Inc.) was used to examine the spatial distribution of calcium channels. The wavelength of the excitation beam was set to 488 and 568 nm (argon ion laser, 4.0 mW). The emitted fluorescence from labelled cells was split by a dichroic mirror (565 nm) into two wavelength bands. The intensities of the green (fDHP) and red (RH414) fluorescent labelling were recorded by two photomultipliers with cut-off filters at 530 and 590 nm, respectively.

To calculate the ratio of fluorescence intensity between green (fDHP) and red (RH414), each confocal image was split into two grey-scale images registering green and red fluorescence intensity. Images were then generated by determining the ratio of the intensity of green fluorescence (fDHP) to that of the red fluorescence (RH414) using the image math function in ‘NIH image software’ (National Institute of Health, USA at http://rsb.info.nih.gov/nih-image/about.html). Only optical sections, 1 μm thick, with sharp and definitive membrane labelling by RH414 (red images) were used for data analysis as this indicated a perpendicular section through the cell membrane. Data were collected and measured using the tool function in NIH image software by drawing lines along the membrane where RH414 showed sharp and definitive labelling. Unless otherwise stated, results are expressed as mean ± s.e.m.

The subumbrellar, muscle sheets of hydromedusae are formed by a monolayer of striated muscle cells. Each cell is polarised with contractile ‘feet’, containing the myofibrils, attached by a narrow neck to the cell soma where the nucleus and other organelles are located (Spencer, 1979). Cell dissociation at room temperature resulted in all cells having their muscle feet retracted with obvious striated myofibrillar bands within the cytoplasm (Fig. 1A), whereas cells dissociated at 30°C preserved their polarised morphology with about 53±5 % of muscle cells having two pairs of muscle feet (Fig. 1B,C). Whole-cell recordings showed that cells dissociated at 30°C not only retained their polarised morphology but also had more active voltage-gated channels as judged by current amplitude (Fig. 1D,E). Of the 87 heat-treated cells we examined, 75 had inward currents, whereas only four of the 20 cells dissociated at room temperature had inward currents. Occasionally some muscle cells that were dissociated at room temperature were tear-shaped with remnants of the muscle feet (Fig. 1A, arrow). All four cells that had some inward current after being dissociated at room temperature were tear-shaped.

In the following experiments all cells were dissociated at 30°C. Calcium currents were isolated by blocking the outward potassium current with Cs²⁺ and TEA applied intracellularly through the patch-pipette. Contamination by sodium currents was checked by using a calcium-free bath solution in which calcium ions and a portion of the NMG-Cl were replaced by 50 mmol l^–1 EDTA-Na₂ or 100 mmol l^–1 NaCl. In both cases Na⁺ concentration was raised to 100 mmol l^–1. We only observed inward current in the calcium-free solution containing EDTA (N=4), not in the absence of EDTA (N=5). This confirmed the presence of calcium channels but not sodium channels. In this case the inward current, in a calcium-free bath solution, was carried by sodium ions passing through calcium channels, while in the absence of EDTA calcium channels were blocked by Mg²⁺ and no inward current was recorded. Fig. 2A shows a typical series of traces of an isolated strontium current. The channels carrying this current activated at approximately –70 mV with maximal current at –30 mV (Fig. 2B, N=10). Current density was calculated to be 0.18±0.05 mA cm^–2 (N=20) by using the maximum current at –30 mV and whole cell capacitance (5.49±1.07 pF; N=20), assuming that membrane capacitance was 1 μF cm^–2. No reversal potential could be detected using test pulses up to +90 mV.

The ionic permeability of calcium channels to barium and strontium was measured (current traces not shown). Channels were equally permeable to barium and strontium with P_Ba/P_Ca=0.86±0.08 and P_Sr/P_Ca=0.85±0.05; N=4 cells in each case.

The rate of activation was voltage-dependent (Fig. 3A) with the time-to-peak for maximal current of 1.49±0.23 ms at –30 mV (N=10). The time course for inactivation was also voltage-dependent (Fig. 3B) with the inactivation time constant varying between 1.5 ms at +80 mV and 3.5 ms at –70 mV (N=10).

Fig. 4A shows the steady-state inactivation curve for Sr²⁺ current. The current was almost fully available at –100 mV, half inactivated at –80±3.5 mV, and completely inactivated at –45 mV. The time constant for recovery from inactivation was 51±0.8 ms (N=10) and complete recovery occurred in 200 ms (Fig. 4B).

We encountered severe channel run-down during recording so that in most cases no current could be recorded 5 min after starting test protocols. Attempts to decrease run-down by addition of ATP and GTP to the electrode solution were not successful. For this reason the selectivity of calcium channel blockers was first examined by evaluating their effect on contraction of muscle strips using field stimulation. When stimulated at 0.2 Hz at 12–14°C in ASW these preparations could maintain a constant contractile force for several hours. Muscle contractile force was decreased in the presence of both dihydropyridines (DHP) and verapamil (Fig. 5). The dihydropyridines nicardipine and nifedipine completely inhibited contraction at concentrations of about 100 μmol l^–1, whereas other dihydropyridines, (+) and (–) Bay K 8644 and nitrendipine, completely inhibited contraction at concentrations less than 40 μmol l^–1 (Fig. 5). Diltiazem at 100 μmol l^–1 and DMSO at 0.5 % had no effect (data not shown).

The effect of dihydropyridines on dissociated cells was also examined to see if blockade of T-type calcium channels could account for the loss of contractility. Only the doses that gave complete inhibition of muscle contraction were used. Fig. 6 shows typical traces of the inhibitory effects of nifedipine at 100 μmol l^–1 (76.08±4.60 %, N=4), nitrendipine at 40 μmol l^–1 (78.02±5.41 %, N=3), (+) Bay K 8644 at 20 μmol l^–1 (81.37±4.56 %, N=5), verapamil at 100 μmol l^–1 (78.22±3.20 %, N=3), and NiCl₂ at 50 μmol l^–1 (57.28±6.00 %, N=4).

Because larger amplitude membrane currents were recorded from dissociated cells that had been heat-treated, and since they also retained muscle feet, we surmised that there was a differential pattern of channel distribution over the cell surface, with channels being preferentially inserted in, or migrating to, the membrane overlying the muscle feet. We used fDHP to label calcium channels since we had determined that these channels were blocked by DHPs. RH414 was used as a reference dye to correct for possible artefacts caused by membrane topology. Fig. 7A shows double-labelling of muscle cells dissociated at room temperature with fDHP (green) and RH414 (red). These cells became spherical, with RH414 labelling only the plasma membrane while fDHP labelled mostly small vesicles within the cytoplasm though occasionally fDHP labelling could be seen on the membrane. Fig. 7B shows double-labelling of muscle cells dissociated at 30°C, which always showed both RH414 and fDHP labelling. There was an obvious association of fDHP labelling with the sarcolemma of the muscle feet. To analyse the differential distribution pattern of fDHP labelling, the ratio of the intensity of green versus red fluorescence of images was measured pixel-by-pixel using NIH imaging software. Larger ratios indicated a higher density of fDHP labelling. The ratio of labelling intensity (fDHP to RH414) in the sarcolemma around the muscle feet was significantly higher than the somatal membrane (0.88±0.03 and 0.67±0.02, respectively, P<0.05, N=20 cells; Fig. 7C).

Our data suggest that heat treatment ‘locks’ the cytoskeleton, thereby ‘fixing’ cell morphology after dissociation. Heat treatment has been used previously during dissociation of tissues from other marine hydrozoan medusae and polyps for the purpose of cell identification (Schmid et al., 1981; Plickert and Kroiher, 1988). Like Schmid and colleagues (Schmid et al., 1981), we noticed that there was a relatively short window of time and temperature for heat ‘fixation’ of the cellular phenotype, and this was particularly the case for retention of electrophysiological competence. Prolonged treatment at 30 °C (>30 min) or treatment at a higher temperature (37 °C) for 15 min preserved cell morphology, but ionic currents could not be recorded. Heat treatment did not apparently alter the kinetics of the inward current, since when compared with cells dissociated at room temperature qualitatively similar kinetics were observed, though current amplitude was markedly reduced (Lin et al., 2000). Inward currents were so small after dissociation at 20°C that statistically relevant data could not be obtained.

Since only cells with intact feet showed large currents, it is possible that most of the channels are located on the feet. Our fDHP data also support the proposition that DHP-sensitive ion channels are at a higher density on the feet. Several other excitable cells also show a differential distribution pattern of ion channels that are obviously related to the excitability properties of the cell in question. For example, voltage-gated sodium channels are known to be clustered at the nodes of Ranvier in vertebrate myelinated axons and neuromuscular junctions (Ritchie and Rogart, 1977; Ellisman and Levinson, 1982; Lupa and Caldwell, 1994), and L-type calcium channels are known to aggregate in T-tubules (Carl et al., 1995). Jellyfish striated muscle cells are morphologically separated into a somal and a ‘foot’ region (Spencer, 1979). These muscle feet contain the contractile myofibres that use extracellular calcium during excitation–contraction coupling (Spencer and Satterlie, 1981). Localisation of calcium channels on muscle feet in jellyfish striated muscle is further supported by the presence of membranous calcium stores just beneath the membrane of muscle feet (Lin and Spencer, 2001). A close association between calcium channels and calcium stores would allow for possible direct electrical coupling or calcium-induced calcium release to augment the influx of calcium through voltage-gated channels for regulation of muscle contraction. The precise way ion-channel proteins and the cytoskeleton interact mechanically is not known, especially for jellyfish. Nevertheless, several adapter proteins have been reported to be associated with voltage and ligand-gated ion channels, which link them to the cytoskeleton. For example, ankyrin and spectrin immobilise sodium channels with actin filaments at presynaptic terminals and along axons (Srinivasan et al., 1988), while rapsyn links nicotinic acetylcholine receptors to actin filaments at neuromuscular junctions (Apel et al., 1995) and gephyrin links glycine receptors to microtubules at postsynaptic sites (Kirsch et al., 1993). GABA_A-receptor-associated protein (GABARAP) links GABA_A receptors to microtubules at synapses (Wang et al., 1999).

The electrophysiological characteristics of the calcium currents recorded from isolated jellyfish muscle cells are similar to those of T-type currents: activation occurred at low voltages (–70 mV), with both activation and inactivation being very rapid and voltage-dependent. Moreover, this current showed another characteristic of T-type channels (Randall and Tsien, 1997) wherein, with progressively stronger depolarisations, successive currents activated and inactivated faster. A noticeable difference between the current we recorded and T-type currents in other systems was the greater rate of both activation and inactivation in jellyfish muscle cells. Time-to-peak for maximum current at a membrane potential of –30 mV was 1.49±0.23 ms and the inactivation time constant was 2.59±0.38 ms for the calcium currents recorded from jellyfish muscles. In comparison the activation time constants for most other T-type currents are usually greater than 10 ms, with inactivation constants greater than 20 ms (Huguenard, 1996; Randall, 1998; Randall and Tsien, 1997).

Our experiments examining the effects of various known calcium channel blockers on muscle contraction were an indirect method for evaluating channel properties, but they also provide us with estimates of the doses that might be effective when using whole-cell, patch-clamp recording. Although dihydropyridines are known to have higher inhibitory potency for L-type calcium channels than T-type channels (Hille, 1992), the sensitivity of the jellyfish calcium current to dihydropyridines, with an IC₅₀ in the μmol l^–1 range, was similar to that reported for T-type calcium channels found in some vertebrate cells, including mouse spermatocytes (Arnoult et al., 1996; Lievano et al., 1996; Santi et al., 1996), hippocampal CA1 neurones (Takahashi and Akaike, 1991), cerebellar Purkinje cells (Kaneda et al., 1990), amygdaloid neurones (Kaneda and Akaike, 1989), rat hypothalamic neurons (Akaike et al., 1989) and mouse sensory neurones (Richard et al., 1991). LVA T-type calcium currents have been reported in some excitable cells of invertebrates (Hagiwara et al., 1975; Deitmer, 1984; Yamoah and Crow, 1994; Ödblom et al., 2000), where the LVA calcium currents in cardiac myocytes of squid are shown to be sensitive to dihydropyridines at micromolar levels (Ödblom et al., 2000). Nickel ions are known to block the T-type calcium channels selectively (Hille, 1992). Jellyfish LVA calcium currents were sensitive to low concentrations of Ni²⁺ (<100 μmol l^–1), which is similar to reports for rat Purkinje cells (Regan, 1991) and amygdaloid neurones (Kaneda and Akaike, 1989). Based upon channel kinetics, activation and inactivation ranges and pharmacological properties, these LVA calcium channels in jellyfish striated muscle cells most closely resemble those of T-type channels.

Due to their low activation threshold and fast inactivation, T-type currents are generally believed to function in combination with other currents to produce pacemaker potentials where rhythmical activity is required (Randall, 1998; Tsien, 1998). In addition, T-type currents are implicated in regulation of cell growth and proliferation (Hermsmeyer, 1998; Triggle, 1998). As the LVA calcium current appears to be the only significant inward current in jellyfish striated muscle cells, this is the first case reported of a LVA calcium current being expressed in isolation without other inward currents. The action potentials of these jellyfish muscles, which have been recorded in current-clamp with sharp electrodes (Spencer, 1978; Spencer and Satterlie, 1981), are reminiscent of those in vertebrate cardiac muscle having large amplitudes (>100 mV) and a long plateau phase (up to 170 ms). Although the ionic basis of the action potential was not clearly established by Spencer and Satterlie, it was demonstrated that most of the inward current was carried by calcium. However, some sodium was required externally to support an action potential. Our results contradict this study; nevertheless it is obvious that the rising phase of the action potential is mostly due to calcium influx through LVA channels, with a delayed repolarisation due to a delayed, rectifier-like potassium current and/or a calcium-dependent potassium current. It must be assumed that the resulting increase in cytoplasmic calcium concentration initiates muscle contraction.

Due to their phylogenetic position at the base of the metazoan radiation, and being one of the first animal groups to possess sodium currents, cnidarians have been a popular taxon in which to look for ancestral types of sodium channels or hybrid calcium/Na⁺ channels (Anderson, 1987; Spafford et al., 1996). Hille (Hille, 1992) noted that T-type calcium channels could be the ancestor of sodium channels, based on the similarity of their kinetics of activation and inactivation. Furthermore, sequence comparisons between sodium and calcium channels show the relatedness of domains in these two channel types (Spafford et al., 1998). Calcium selectivity of HVA channels is conferred by four negatively charged glutamate residues, which form a high-affinity, EDTA-like calcium-binding site in the pore region (Tsien, 1998). This contrasts with sodium channels where two of the negatively charged residues are replaced by a positively charged residue (lysine) and a neutral residue (alanine) (Heinemann et al., 1992; Ellinor et al., 1995). T-type channels might represent the transition state where these four critical residues are two glutamates and two aspartates (Tsien, 1998). Measurements of sequence similarity between L-type, non-L-type, T-type and sodium channels also suggests that there was an early divergence of these channel types (Tsien, 1998). However, the presence of a functionally undefined, glycosylation-rich, extracellular loop between segments 5 and 6 (located just in front of the pore in domain I and found only in sodium channels and T-type calcium channels) might represent a remnant of their shared ancestry (Spafford et al., 1998).

This jellyfish LVA current activates and inactivates much faster than most vertebrate T-type currents reported to date, with kinetic properties similar to sodium currents. In jellyfish striated muscle cells this calcium current is the major inward current of action potentials, and it is tempting to speculate that this LVA calcium channel is closely related to an ancestral sodium channel.

This article is dedicated to Nikita Grigoriev, a scientist with a technical acumen that, in our experience, was unsurpassed. We thank the staff at Bamfield Marine Station for assistance in collecting jellyfish and Mr R. Bhatnagar and Mr R. Mandryk for technical support. This study was supported by a grant to A.N.S. from the Natural Sciences and Engineering Research Council, Canada.