Na⁺, K⁺ and Ca²⁺ Currents in Identified Leech Neurones in Culture

Detailed information is available about the properties of identified leech neurones. For many nerve cells of known function, the morphology, the transmitter chemistry and the synaptic connections have been studied within the CNS; from cell type to cell type there are striking differences in action potential configuration, delayed rectification, long-lasting Ca²⁺-activated K⁺ conductances and other variables that influence signalling. Each cell has an ensemble of properties by which it can be unambiguously recognized (Nicholls, 1987). Moreover, after an identified neurone has been removed from the ganglion and placed in culture, it retains these properties. For example, sensory cells continue to give highly distinctive action potentials for days or weeks in culture.

At the same time, surprisingly little is known about how the various identified nerve cells differ with respect to the ion channels they exhibit. From experiments made on neurones within ganglia, it is known that action potentials in leech neurones depend on Na⁺, are unaffected by moderate concentrations of tetrodotoxin (Nicholls & Kuffler, 1964; Kleinhaus & Prichard, 1983; Beleslin, 1985) and are prolonged by TEA⁺ and 4-aminopyridine (Kleinhaus, 1976; Kleinhaus & Prichard, 1975; Johansen & Kleinhaus, 1985,1986). However, numerous technical difficulties arise if one wishes to compare with voltage-clamp the ionic currents that give rise to the membrane properties in different types of cells. These include: (1) the large sizes of the cells which makes conventional whole-cell patch-clamp difficult to use; (2) the high resistances of the microelectrodes necessary for penetrating leech neurones when using two-electrode voltage-clamp; (3) the complex arborizations of the cells in situ which prevent adequate space clamping; and (4) the ineffectiveness of tetrodotoxin.

In this paper we describe voltage-dependent Na⁺, K⁺ and Ca²⁺ currents in Retzius (R), anterior pagoda (AP) and sensory [pressure (P), nociceptive (N) and touch (T)] cells in culture. A particular advantage of the cells in culture is that they can be effectively voltage-clamped because they are isopotential spheres when plated on polylysine, a substrate that does not promote process outgrowth (Chiquet & Acklin, 1986; Dietzel et al. 1986; Ross et al. 1987). Emphasis has been placed on R and AP cells, which display markedly different K⁺ currents, and on P sensory cells with which R cells form chemical synapses in culture (Fuchs et al. 1982). Divalent cation currents are described in more detail for R cells elsewhere (R. J. Bookman & Y. Liu, in preparation). These experiments constitute a necessary first step in the analysis of currents responsible for generating the distinctive action potentials of identified cells. Moreover, information about manipulations required for blockage of Na⁺ and K⁺ currents will make it possible to correlate inward Ca²⁺ currents with transmitter release at synapses formed in culture by R and P cells.

The methods for removing and culturing leech neurones have already been described (Fuchs et al. 1981 ; Dietzel et al. 1986). R, AP, T, N and P cells identified by their sizes and positions were removed from desheathed ganglia by lassoing them with nylon monofilament loops or by suction after treatment for 1 h at room temperature with collagenase and dispase (2 mg ml⁻¹; Boehringer-Mannheim Corporation). The diameters of these neurones were, in order from largest to smallest, R (80–100 μm) > AP > P > N > T (50 μm). Individual cells were placed in Leibowitz-15 (L-15, Gibco) medium supplemented with 0·1 mg ml⁻¹ gentamycin, 6 mg ml⁻¹ glucose and 2 % foetal calf serum and plated onto polylysine-coated microwells (Falcon 3034F, type 60) for 1– 14 days at 20°C.

Two-electrode voltage-clamp (Almost Perfect Electronics, Basel, Switzerland) was used to measure the voltage-dependent ionic currents in each cell type (see Fig. 1A). Approximately 75% of the cells penetrated yielded useful results. All measurements were made at room temperature (22–25 °C). Cells were observed in an inverted microscope. Electrodes were pulled from Haer 30–30–0 thin-walled microelectrode glass and filled with 4 mol l⁻¹ potassium acetate. 10–20 MΩ electrodes were used for passing current and 10–30 MΩ electrodes for measuring voltage. The voltage was recorded differentially (see Fig. 1A) and was connected to the feedback circuit of the voltage-clamp. A large, grounded aluminium shield was interposed between the electrodes to reduce capacitative coupling between them. Owing to the high resistance of the electrodes that are needed to penetrate leech neurones, a number of special precautions were necessary to ensure adequate clamping and short time constants. The current signal from the voltageclamp was fed through a high-frequency amplifying circuit (Almost Perfect Electronics, Basel, Switzerland) to compensate for the time constants of the two electrodes and was connected to the current input of a second preamplifier. This, in turn, was connected to a high-voltage, constant-current head stage (± 100V) which was used to pass current through the current-passing microelectrode. Current and voltage signals were stored on magnetic tape and transferred to a computer (MINC/DECLAB-23) for analysis as described below.

For measuring the total current evoked by depolarizing voltage steps, the external solution was standard leech Ringer’s fluid to which A-methyl-D-glucamine chloride was added to bring the osmolality to between 350 and 370 mmol kg⁻¹. This corresponds to the tonicity of the L-15-enriched solutions in which the neurones were cultured. The solutions used for measuring individual currents are listed in Table 1.

A gravity-fed perfusion system was used to exchange bathing solutions. The solution was fed into the microwell through a pipette and the height of the solution reservoir as well as the diameter of the tip of the pipette were adjusted to regulate the flow rate. Typically, a complete change of solution, as judged by changes in ionic currents, was achieved in 2–3 min.

After transfer to a computer, current records produced by depolarizing voltage pulses had their capacitative transients and leak currents subtracted. The current records used for subtraction were produced either by a hyperpolarizing pulse (p) of equal magnitude to the depolarizing pulse or by three hyperpolarizing pulses (p/3) each one-third the magnitude of the depolarizing pulse. Holding currents were then subtracted to give a zero-baseline before further analysis. The activation time (t_1/2, time to one-half peak amplitude) and inactivation time constant (τ_h) of the current were determined. In some experiments, a family of current and voltage traces was used to generate peak current–voltage curves. In others, the effect of a blocking agent was monitored by measuring against time the current produced by pulses to a particular voltage.

Statistical comparison was made using Student’s t-test, and differences were considered significant at P<0·05.

In saline containing normal or half-normal [Na⁺] (solutions 1 and 2, Table 1) step depolarizations of the membrane potential (from −50 mV to between −30 and +10 mV) produced a rapid inward current followed by a delayed outward current.

Hyperpolarization of the membrane produced only capacitative artefacts at the onset and offset of the pulse and a small, sustained leakage current during the pulse. Examples of these currents in an R cell are shown in Fig. 1B,C. The following questions have been examined. (1) Which ions carry these currents? (2) How can each current be separated from the others? (3) How do the separate ionic currents differ in the various cell types?

In leech neurones, although external Na⁺ is necessary for action potentials, moderate concentrations of tetrodotoxin or saxitoxin have no effect (Beleslin, 1985; Johansen & Kleinhaus, 1987). To determine whether Na⁺ was responsible for the initial inward current, the external Na⁺ concentration was lowered from 125 to 10 mmol l⁻¹ while applying depolarizing steps to −5 mV (solutions 1 and 4, Table 1). In all five types of cell (R, AP, P, N and T) as the concentration of Na⁺ decreased during the exchange of solutions the inward current decreased and eventually reversed. After replacing normal-Na⁺ saline the inward current reappeared (Fig. 2).

Activation kinetics for the Na⁺ current were too fast to be measured by conventional two-electrode voltage-clamp, owing to the large capacitance artefacts (see Materials and methods). They are described elsewhere (R. J. Bookman, J. G. Nicholls, H. Reuter & W. B. Adams, in preparation). Inactivation kinetics could, however, be analysed after reduction of Na⁺ to 50 % of normal concentration (which allowed for better control of membrane voltage during the peak Na⁺ current), while Mn²⁺ and TEA⁺ blocked Ca²⁺ and K⁺ currents (solution 3, Table 1; Fig. 3A). There were no significant differences between inactivation time constants of Na⁺ currents in the different types of cultured cells. Fig. 3B shows time constants of inactivation in P and R cells with graded voltage steps from a holding potential of −50 mV. For voltage steps from −11 to −1 mV, Na⁺ currents were sufficiently large and slow to be measured accurately.

In contrast to the similarity between Na⁺ inactivation kinetics in different cell types, the maximum peak Na⁺ current and the current density showed marked differences. The membrane surface areas of R and P cells were estimated from measurements of membrane capacitance derived from input resistance and time constant. For R cells the values were: 2·0 ± 0·2 nF and 16 ± 4 MΩ (mean and S.E., N = 7); for P cells the corresponding values were: 0·9± 0·4 nF and 15 ± 4MΩ (mean and S.E., N=7). The mean maximum peak Na⁺ current density calculated in this way for R cells was 11 ± 2 μAcm⁻² (mean and S.E., seven cells) and for P cells was 53 ± 11 μAcm⁻² (mean and S.E., seven cells). The difference between the means was highly significant (P< 0·005). Not surprisingly, Na⁺ current density was greatest in cells that produced the largest action potentials, and was smallest in AP cells.

Neurones bathed in low-Na⁺, TEA⁺ medium containing 10 mmol l⁻¹ Ca²⁺, Ba²⁺ or Sr^24- exhibited inward currents upon step depolarization to between −5 and +30 mV. Examples of inward Ca²⁺ currents in AP, R, P and N cells are shown in Figs 4 and 6A (solutions 5 and 12, Table 1). In all experiments, these inward currents activated and inactivated more slowly than Na⁺ currents and were unaffected by changes in external [Na⁺], Cd²⁺ and Mn²⁺ blocked the divalent cation current reversibly. A slow decline in inward Ca²⁺ current was observed in R, N and P cells, but not in AP cells (Figs 4, 6A). Whether this represents inactivation of Ca²⁺ channels or the result of slow, unblocked outward currents or a combination of both would not be resolved by our techniques. Further analysis of divalent cation currents in R cells is presented elsewhere (R. J. Bookman & Y. Liu, in preparation).

Experiments were performed which demonstrated that the delayed outward current shown in Fig. 1 was carried by K⁺. The external K⁺ concentration was increased to change EK, the equilibrium potential. In leech neurones it has been shown that E_K changes from −85 to −27 mV when the external K⁺ concentration is increased from 4 to 40 mmol l⁻¹ (Nicholls & Kuffler, 1964). In 40 mmol l⁻¹ K⁺ solution, during a step depolarization from −50 to +10 mV, the driving force for K⁺ is reduced. Upon repolarization the K⁺ tail currents should be inward, since E_K is now positive with respect to the holding potential of −50 mV. As expected, increasing external [K⁺] from 4 to 40 mmol l⁻¹ reduced reversibly the peak outward current by nearly half (Fig. 5) and produced inward K⁺ tail currents during repolarization to the holding potential of −50 mV.

Additional evidence for the role of K⁺ was provided by reversibly blocking outward currents with 25–100 mmol l⁻¹ TEA⁺ (solution 5), with 0-1-1 mmoll^-1 quinidine (added to solution 4) and with 5–10 mmol l⁻¹ 4-AP (solution 6, Table 1). Fig. 6 shows the effects of 30 mmol l⁻¹ TEA⁺ on outward current in an R cell (solution 5, Table 1). Similar blockade of outward currents by TEA⁺ and quinidine was seen in AP, T, P and N cells. 4-AP reduced K⁺ currents in all five types of cells.

As shown in Fig. 7, the outward K⁺ currents in AP, R, P and T cells differed markedly. The currents were larger and faster in AP and R cells than in the sensory cells. To categorize the various K⁺ currents the following measurements were made in each type of cell: (1) the rate of activation, which was not a simple exponential, was characterized by the t_1/2 to peak; (2) inactivation was described by its time constant and by the effect of various levels of steady holding potential; (3) Ca²⁺-activated K⁺ currents were identified by applying Ca²⁺ channel blockers.

Four types of K⁺ current were distinguished; they have been designated as I_A (rapid activation and inactivation), I_C (Ca²⁺-dependent), I_K1 and I_K2 (analogous to delayed rectifier K⁺ currents of squid axon), in accordance with nomenclature established by others (Adams & Benson, 1985; Kaczmarek & Levitan, 1987). The currents are described for each type of cell in the following paragraphs and summarized in Table 2. Our principal emphasis has been to establish the distribution of the various currents in different cells rather than to define their kinetics and voltage-sensitivity in detail.

Complex, intermingled K⁺ currents with markedly different characteristics were apparent in R cells. It was of importance for studies in which R cells make synapses onto P cells and release 5-hydroxytryptamine (5-HT) to establish the identities of these currents and to determine how they could be selectively eliminated by pharmacological agents or by inactivation.

A major K⁺ current was Ca²⁺-activated and corresponded to I_C. Thus, the outward current was reduced by 64 ± 5% (S.E.M., N =6) in the presence of 100 μmol l⁻¹ Cd²⁺, or after substitution of Mn²⁺ for Ca²⁺ (solutions 7, 11 and 14, Table 1; Fig. 8A). Fig. 8B shows the N-shaped current-voltage relationship of K⁺ currents in fluid containing 2 mmol l⁻¹ Ca²⁺. After addition of 100 μmol l⁻¹ Cd²⁺, the outward current changed to a monotonic function with increasing steps of depolarization. The separation of K⁺ currents in R cells, into Ca²⁺-activated and Ca²⁺-independent, was further supported by the measurements of activation shown in Fig. 8C. Again after addition of 100 μmol l⁻¹ Cd²⁺, the time to reach half-peak current was a simple monotonic function, activation becoming faster with larger depolarizing steps. In contrast, the relationship was complex under conditions allowing Ca²⁺ entry and activation of I_C. The time to half-peak decreased with steps of depolarization up to about +30 mV and then increased, presumably reflecting changes in the driving force on Ca²⁺. I_C in R cells was not blocked by high concentrations of charybdotoxin (95 nmol l⁻¹) or apamin (10 μmol l⁻¹) which are peptide blockers of other Ca²⁺-activated K⁺ channels (Hugues et al. 1982; Miller et al. 1985).

At least three other K⁺ currents with distinctive features could be recognized in R cells. As shown in Fig. 8, a K⁺ current, designated I_KX, remained after blockage of I_C; IKI was slower than I_C and was blocked in the presence of 25–100 mmol l⁻¹ TEA⁺ (solutions 4 and 5, Table 1). A second, even slower current, IK2, ^was observed during very long depolarizing pulses (1–2 s) in the presence or absence of I_c (Fig. 9). Increased external [K⁺] (from 4 to 23 mmol l⁻¹) reduced peak I_K1 and I_K2 currents as expected, and reversed the tail currents observed upon repolarization to −50 mV. K⁺ currents similar to I_KI and I_K2 predominated in sensory T, P and N cells (see below).

In addition to I_C, I_K1 and I_K2, a rapidly activating and inactivating A-type current (I_A) was revealed in R cells. This residual A-current was present after blockage of I_C, I_KI and I_K2 (by low-Na⁺, Ca²⁺-free, Mn²⁺, TEA⁺ fluid; solution 15, Table 1) and was reduced further by addition of 5 mmol l⁻¹ 4-AP. It is described in greater detail for AP cells, below. Unlike the slower K⁺ currents, I_A was completely inactivated at holding potentials more positive than −40 mV (see Fig. 10C).

The predominant K⁺ current in AP cells was of the A-type (I_A). Blockage of Ca²⁺ entry did not decrease outward K⁺ currents in AP cells as it did in R cells. Instead, a small increase in outward current was observed, presumably owing to the abolition of a competing inward Ca²⁺ current. Thus Ic appears to be absent from AP cells. Nor did we find any trace of outward currents that activated and inactivated slowly with kinetics similar to those of I_K1 and I_K2

Although activation of I_A in AP cells was rapid and similar to activation of outward K⁺ current in R cells (Fig. 7B), inactivation of I_A was considerably faster. The time constant for inactivation (r_h) for AP cells was 26 ± 2 ms (S.E.M., N= 8) compared with 97 ± 12ms (S.E.M., N = 10) for R cells, when tests were made with prolonged steps from −60 to 0 mV.

I_A also showed pronounced steady-state inactivation. When the holding membrane potential was changed from −70 to −30mV in an AP cell, the K⁺ current evoked by a step to 0 mV was reduced by 81 ± 6 % (S.E.M., N = 6). The absence of I_C, I_KI and I_K2 currents in AP cells suggested that the small outward current that remained when holding at −30 mV represented I_A that had not been completely inactivated. It was, however, slightly slower in its activation than the major outward A-type current. Thus, for residual I_A t_1/2 of activation starting from −30mV was 3·2 ± 0·5 ms (S.E.M., N = 4) compared with t_1/2 starting from −70mV of 2·1 ± 0·18 ms (S.E.M., N = 6) for total I_A (P<0·05). This slowing of activation appeared to be due to a competing inward Ca²⁺ current (see example of Ca²⁺ current in Fig. 4).

A comparison was made of the effects of membrane holding potentials on residual I_A of AP and R cells in solutions containing 100 mmol l⁻¹ TEA⁺ and 10 mmol l⁻¹ Mn²⁺ (Fig. 10C; solution 15, Table 1). The amplitude of I_A in both AP and R cells behaved similarly and declined steeply at holding potentials between −50 and −35 mV.

The K⁺ currents in P, N and T cells were slower than those in AP and R cells (Fig. 7) and resembled the delayed rectifiers described for squid axons, designated as I_Ki (intermediate) and I_K2 (slower activation). I_K1 predominated and in certain sensory neurones was the only detectable K⁺ current. To measure activation and inactivation, depolarizing pulses were applied after the membrane potential had been held at various steady levels (from −70 to −40 mV). In P cells the half-time to reach peak (t_1/2) was 16 ± 2ms (S.E.M., N = 7) for voltage steps to −10mV. With larger steps of depolarization (to +35 mV) the half-time of activation decreased to 6ms (N = 3; see Fig. 7B). Inactivation of I_K1 in P cells was slow and followed an exponential time course (τ_h = 423 ± 58 ms; S.E.M., N = 15).

In N cells, activation and inactivation were even slower for voltage steps to 0 mV (t_1/2 = 31 ± 5 ms, S.E. M., N = 5; τ_h = 1700 ± 280ms; S.E.M., N = 7). T cells had the fastest activation of sensory neurones for voltage steps to 0 mV (t_1/2 = 9ms, N = 4); inactivation, however, was similar to that of P cells (τ_h = 402 ms, N = 3).

For reasons that are not clear, I_K2, the slowly activating K⁺ current, was evident in only certain of the cultured neurones. I_K2 was characterized by its very slow activation kinetics which caused the outward current to rise continually during a maintained pulse without reaching a plateau in 2 s. This was the longest pulse that could be applied without damaging the cell irreversibly. The slow progressive creep of I_K2 activation was still present in raised external [K⁺] (23 mmol l⁻¹), showing that it was not caused by K⁺ accumulation.

In spite of the wealth of information about the electrical properties and signalling characteristics of identified neurones in the CNS of the leech Hirudo medicinalis, there has till now been no attempt to describe the major cationic currents in the various cell types. An exception is the analysis of K⁺ currents with two-electrode voltage-clamp in Retzius cells in the CNS of a different leech, Macrobdella decora (Johansen & Kleinhaus, 1986). Technical difficulties have stood in the way of more comprehensive analyses of the distribution and the characteristics of Na⁺, Ca²⁺ and K⁺ currents in leech neurones. First is the absence of a selective Na⁺ channel blocker; instead of simply applying tetrodotoxin, the external Na⁺ concentration must be lowered. Worse still are the relatively small sizes of the cells which make it necessary to impale them with two fine, high-resistance microelectrodes. As a result it becomes difficult to deliver sufficient current to control the membrane potential and the capacitative artefacts can encroach upon the early phases of activation, especially for Na⁺.

With such considerations in mind, the aims of the present experiments were limited. The use of an isolated neurone in culture offers the advantage that one can, with confidence, clamp the cell in its entirety and change its fluid environment rapidly. Conventional two-electrode voltage-clamp seemed suitable as a first step in comparing channel types and densities in those neurones most frequently studied in culture. The results presented here were not designed to measure precise kinetics of Na⁺, K⁺ and Ca²⁺ activation and inactivation, for which other techniques such as patch-clamp, intracellular perfusion or loose-patch (Bookman & Dagan, 1987; R. J. Bookman & Y. Liu, in preparation; Bookman et al. 1987) can offer better temporal and spatial resolution. Rather, the aim was to establish whether different types of channels for cations existed in R, AP, P, N and T cells. These cells were selected because they show highly distinctive action potentials, delayed rectification and afterpotentials that are retained in culture. Moreover, in culture, extensive use has been made of R and AP cells to study neurite outgrowth and of R and P cells to study synapse formation (Nicholls, 1987).

Our results lead to certain clear conclusions. The time constants for inactivation of Na⁺ currents (unlike the activation kinetics) could be measured accurately, as could maximum peak Na⁺ current. It became evident that P and N cells, with their large action potentials, did not differ from R, T and AP cells with respect to Na⁺ inactivation but only in the density of Na⁺ channels. Similarly, what distinguished the Ca²⁺ currents from cell to cell was the peak amplitude rather than the rate of rise. For K⁺ currents the picture was more complex and remains incomplete. Simplest were the AP cells with the fastest activating and inactivating K⁺ currents. This, together with the minuscule Na⁺ currents, can account for the small amplitude of impulses in the soma, as well as the high rates of firing and the absence of delayed rectification that together identify the AP cell electrically.

The analysis is less satifactory for the sensory cells. For convenience and by analogy we have labelled the K⁺ currents as I_K1 and I_K2. These currents were far slower and could help to explain the larger sizes of action potentials, the undershoot, and the delayed rectification in T, P and N cells. Activation of I_K1 was fastest in T cells, followed by P and then N cells. These differences in activation probably correlate with the rapid repolarization of action potentials of T cells compared with the slow repolarization of N cells. But we cannot account for these phenomena quantitatively and numerous other anomalies remain. For example, no evidence for Ca²⁺-activated K⁺ current was detected in P or N cells; yet these cells show a persistent hyperpolarization that depends on Ca²⁺ and long-lasting changes in G_K following trains of impulses (Jansen & Nicholls, 1973). A peculiarity that may be related concerns the very slow activation of I_K2 which was seen in some P and N cells, but not others, in a manner not correlated with other variables.

Currents resembling I_K1 and I_K2 of leech sensory cells have been found in various different proportions in neurones of other invertebrates (e.g. see fig. 13 of Aldrich et al. 1979). One mechanism for such variability is thought to be phosphorylation of delayed rectifier K⁺ channels through second-messenger systems. For example, increasing cyclic AMP levels in bag cells of Aplysia partially eliminates the fast-activating delayed rectifier K⁺ current and completely eliminates the slowly activating K⁺ current (Strong & Kaczmarek, 1986). It remains to be determined whether comparable K⁺ channel regulation also occurs in leech neurones.

Two pairs of homologous N cells exist in each midbody ganglion of Hirudo medicinalis: a lateral and a medial pair. Lateral and medial N cells are similar in their functions and action potential shapes, but are different in that each type of cell exhibits specific antigens (Johansen et al. 1984a; Zipser, 1982; Blackshaw, 1982) and extrasynaptic receptors (Sargent, 1977; Johansen et al. 1984b). In our experiments we used N, T and P cells without taking account of subtypes. It could therefore be that analysis of K⁺ currents in lateral and medial types of N or P cells might show subtle differences.

R cell K⁺ currents were more complex and their functional significance harder to interpret. Not only were all the K⁺ currents of the other types of cells displayed but there was an additional major Ca²⁺-activated component. To what extent I_c, IA, IKI and IK2 currents represent different populations of channels with distinctive characteristics could not be resolved in our experiments in which separation of the component K⁺ currents was achieved by drastic changes in ionic, TEA⁺ and 4-AP concentrations.

At the same time these measurements set the stage for more detailed studies of synaptic transmission by R cells in culture. Synapses made by R cells upon P cells develop rapidly and show properties such as facilitation, depression and dramatic effects of steady holding potential on the release of 5-HT (the Retzius cell transmitter). It now becomes practicable to measure, under conditions defined by the present experiments, Ca²⁺ currents that are relatively uncontaminated by I_A, Ic or Na⁺ currents, and to correlate Ca²⁺ entry with transmitter release during facilitation.

We thank Dr R. J. Bookman for his help and encouragement during each phase of these experiments and Mss H. Niederer and P. Muller for their skilful preparation of cell cultures. We are grateful to Ms J. Wittker for her excellent secretarial assistance and to Mr P. Baettig for photographing the figures. This work was supported by grants from the Swiss National Fund and the US Navy to JGN; RRS was supported by an NIH Postdoctoral Fellowship.