Characterization of Voltage- and Ca²⁺-Activated K⁺ Channels in Squid Olfactory Receptor Neurons

The squid olfactory organ is a small (0.5–1.0 mm diameter), knob-like, ciliated structure located ventrally and posteriorly to each eye near the opening to the mantle cavity. The pseudostratified sensory epithelium contains ciliated support cells, mucus-producing goblet cells, basal cells and several morphological types of olfactory receptor neurons (ORNs) (Emery, 1975). The cell type used in the present study is the pyriform receptor cell also described as the type 4 cell by Emery (1975).

In general, squid ORNs are primary bipolar sensory neurons with an apical, ciliated dendritic region, a soma and a basal unmyelinated axon. The unmyelinated axons from ORNs form bundles that leave the olfactory organ and, as the olfactory nerve, travel through a foramen in the cartilaginous skull to their first synapse in the olfactory lobe in the brain. Further connections are made in the optic gland, which influences gonadal maturation, in the magnocellular lobe, which controls escape jetting, and in the palliovisceral lobe, which is thought to control the emission of ink (Young, 1976). Behavioral studies have shown that the olfactory organ on the head of the squid is the site where chemical sensitivity is highest (excluding the tentacles) and suggest that input from the olfactory organ influences motor centers that control escape jetting (Gilly and Lucero, 1992). K⁺ channel blockers, as well as squid ink, which contains L-Dopa and dopamine (Lucero et al. 1994), were found to activate escape jetting when focally applied to the squid olfactory organ (Gilly and Lucero, 1992). Isolated ORNs from the squid Loligo opalescens have voltage-gated Na⁺, Ca²⁺ and K⁺ currents, and respond to ink, L-Dopa and dopamine with a hyperpolarization that inhibits the firing of action potentials (Lucero et al. 1992). These odor-induced hyperpolarizations could be due to activation of Cl⁻ or K⁺ channels. To help elucidate the role(s) that K⁺ channels play in the transduction of chemical signals to electrical impulses in squid ORNs, we have characterized the voltage- and Ca²⁺-gated K⁺ currents. We have identified three types of K⁺ currents in these cells: a delayed rectifier K⁺ current (I_K), a rapidly inactivating A-type K⁺ current (I_A) and a Ca²⁺-activated K⁺ current (I_K(Ca)). A knowledge of the kinetics, selectivity, voltage-dependence and pharmacology of squid ORN K⁺ currents will be useful for manipulating ionic conditions to study receptor currents in isolation, for modelling the amplitude and frequency output of the cell in response to depolarizing or hyperpolarizing inputs, and for making comparisons with K⁺ channels in other olfactory models. These studies will contribute to our overall understanding of the role of K⁺ channels and hyperpolarizing influences in the coding of odorant information at the level of the single cell.

The dissociation and culture of olfactory receptor neurons was essentially the same as described in Lucero et al. (1992). Juvenile squid (Lolliguncula brevis, Berry; 4–5 cm mantle length) were obtained from the National Resource Center for Cephalopods, Galveston, TX, USA. The animals were killed by decapitation. The olfactory organs were excised and treated with non-specific protease (10 mg ml⁻¹ Sigma type XIV) in sterile filtered artificial sea water (ASW) (see Solutions) at room temperature (20–23 °C) for 50 min. Following a 3–5 min rinse in fresh filtered ASW, the isolated olfactory organ was transferred onto a 12 mm round glass coverslip coated with Concanavalin A (Con A), covered with a drop of sterile culture medium and teased apart using glass needles. The cells were allowed to settle for 10 min before adding 2 ml of culture medium. Dishes of cells were placed in a humidified incubator at 20 °C for periods ranging from 1 h to 3 days before use.

Culture medium consisted of Leibovitz’s L-15 (Gibco Co., Grand Island, NY, USA) supplemented with 163 mmol l⁻¹ NaCl, 3.5 mmol l⁻¹ KCl, 9 mmol l⁻¹ CaCl₂, 35 mmol l⁻¹ MgCl₂, 2 mmol l⁻¹ Hepes (pH 7.6), 2 mmol l⁻¹ L-glutamine, 50 i.u. ml⁻¹ penicillin G and 0.5 mg ml⁻¹ streptomycin (780 mosmol l⁻¹). Artificial sea water (ASW) consisted of 340 mmol l⁻¹ NaCl, 10 mmol l⁻¹ KCl, 35 mmol l⁻¹ MgCl₂, 10 mmol l⁻¹ CaCl₂, 10 mmol l⁻¹ Hepes, 10 mmol l⁻¹ glucose, pH 7.4. Solutions used in testing the relative permeability of the K⁺ channels to Na⁺versus K⁺ were made by equimolar replacements of NaCl with KCl. The 200-K⁺ external solution consisted of 200 mmol l⁻¹ KCl, 140 mmol l⁻¹ choline chloride, 35 mmol l⁻¹ MgCl₂, 10 mmol l⁻¹ CaCl₂, 10 mmol l⁻¹ Hepes, 15 mmol l⁻¹ glucose, pH 7.4. Solutions used for monovalent cation selectivity studies were made by replacing the 200 mmol l⁻¹ KCl with 200 mmol l⁻¹ of the test ions: Na⁺, Rb⁺, Cs⁺ or NH₄⁺. The internal solution for all of the experiments consisted of 20 mmol l⁻¹ KCl, 155 mmol l⁻¹ potassium glutamate, 25 mmol l⁻¹ KF, 1.5 mmol l⁻¹ EGTA, 0.5 mmol l⁻¹ CaCl₂, 10 mmol l⁻¹ Hepes, 70 mmol l⁻¹ glucose, pH 7.2. The liquid junction potentials between the pipette and bath solutions were measured, and reversal potentials for selectivity studies were corrected a posteriori according to Neher (1995). The osmolarity of both internal and external solutions (780 mosmol l⁻¹) was chosen because it is in the mid-range of the osmolarities at which these brackish-water squid (Lolliguncula brevis) live. Unless indicated, chemicals were obtained from Sigma Chemical Co. (St Louis, MO, USA), except for charybdotoxin (CTX) which was a generous gift from NPS Pharmaceuticals, Inc. (Salt Lake City, UT, USA).

Experiments were conducted in a temperature-controlled chamber (12 mm diameter × 2 mm depth). Pharmacological agents were applied using either a loop injector (Rainin Inc., Woburn, MA, USA) or an SF-77 rapid solution changer (Warner Instrument Corp., Hamden, CT, USA), with the exception of CTX which was applied with a Picospritzer II (General Valve Corp., Fairfield, NJ, USA).

Coverslips with adherent cells were placed into the recording chamber and perfused with the external bath solution (ASW) at a rate of 1–2 ml min⁻¹. With the exception of the Q₁₀ measurements, experiments were conducted at 10 °C in order to slow channel kinetics. Patch electrodes with resistances of 1–2 MΩ were pulled from thick-walled (0.64 mm) borosilicate glass on a Flaming/Brown P87 microelectrode puller (Sutter Instrument Co., Novato, CA, USA). The bath reference electrode was a silver/silver chloride pellet placed downstream of the cells. The whole-cell recording configuration (Hamill et al. 1981) was achieved by pressing the patch electrode against the cell while monitoring resistance. Once a gigaohm seal had formed, the holding potential was set to −80 mV, and gentle suction was applied to rupture the membrane under the pipette. We found that −80 mV was the most negative holding potential that could be used for extended recordings.

Voltage-clamp experiments were run using a Digidata 1200 Interface (Axon Instruments, Inc., Foster City, CA, USA) and a 486-33 IBM-clone computer. Membrane currents were measured using an Axopatch 200A integrating patch-clamp amplifier (Axon Instruments, Inc.). Sampling rates ranged from 500 Hz to 100 kHz, and filter cut-offs for the low-pass eight-pole bessel filter ranged from 200 Hz to 10 kHz. PClamp software (Axon Instruments, Inc.) was used to generate pulse protocols and acquire data. Analyses were performed using Webfoot software (Biodiversity, Salt Lake City, UT, USA). A standard leak pulse protocol that consisted of averaging the currents (I) from 24 repeated 12 ms voltage (V) pulses that stepped from −70 mV to −80 mV was used to calculate membrane input resistance (RM), by dividing ΔV by ΔI according to Ohm’s law. Cell capacitances (C_M) were determined by integrating the area under the capacity transient to obtain the amount of charge (Q) moved, and dividing Q by ΔV. A double exponential plot was fitted to the decay of the capacity transient because there are both fast (τ₁) and slow (τ₂) time constants associated with charging the membrane of the soma and cilia of these cells (Lucero et al. 1992). The time constants and current amplitudes from the double exponential fit were used to calculate the resistances in series with the pipette and soma (R_S1) and the cilia (R_S2) according to the method described by Llano et al. (1991) for complex cells. Positive feedback compensated for 80–90 % of the total series resistance (R_S). Linear leak currents and the small residual, uncompensated capacity currents were subtracted from all data on-line using a pre-pulse (P/4) procedure (Armstrong and Bezanilla, 1974). Results are presented as mean ± S.E.M., N=number of cells.

We measured the passive properties of isolated pyriform ORNs from the squid Lolliguncula brevis using the capacitance pulse protocol described in Materials and methods. The mean R_M of 145±8 MΩ (N=30) was lower than that reported for the squid Loligo opalescens (Lucero et al. 1992) and much lower than the input resistance of several gigaohms reported for ORNs from various species (frog, Rana pipiens, Pun and Gesteland, 1991; rat, Trombley and Westbrook, 1991; zebrafish, Danio rerio, Corotto et al. 1996). The mean values for the fast and slow time constants of the capacity transient of τ₁=60±5 μs and τ₂=2.9±0.3 ms (N=26) were similar to those for Loligo opalescens (Lucero et al. 1992). The mean values for the capacitances associated with the somal (C₁) and ciliary (C₂) regions of these cells were 9.0±0.5 pF for C₁ and 63.3±5.4 pF for C₂ (N=26). The uncompensated mean value for the R_S1 associated with the pipette and soma was 8.2±0.4 MΩ, while the equivalent R_S2 associated with access to the ciliary region was 40.2±3.7 MΩ (N=26). The resting membrane potential was measured under current-clamp conditions and averaged −57.5±1.0 mV (N=30).

Whole-cell voltage-clamped squid ORNs have both inward and outward currents. We determined that the outward currents were selective for K⁺ over Na⁺ by replacing external Na⁺ with equimolar amounts of K⁺ in the presence of 200 nmol l⁻¹ tetrodotoxin (TTX). Fig. 1A–D shows the appearance of inward K⁺ currents as external K⁺ concentration ([K⁺]_o) was increased from 10 to 350 mmol l⁻¹. Even in symmetrical 200 mmol l⁻¹ KCl, outward rectification is visible because fewer K⁺ channels are open at the more negative potentials (Fig. 1C). A tails voltage protocol, which steps to +40 mV to open K⁺ channels and then steps to the voltages indicated in the inset of Fig. 1E to change the driving force on the permeant ions and elicit tail currents through the open channels, was used to determine the instantaneous current–voltage (I–V) relationship and reversal potential for each [K⁺]_o. The slope of the mean reversal potential measured for each [K⁺]_o plotted against the log of the K⁺ concentration (in mmol l⁻¹) was 57.6±1.9 mV per decade (N=4; solid line in Fig. 1F). This value agrees closely with the calculated Nernst potential for highly selective K⁺ channels of 56.2 mV per decade at the experimental temperature of 10 °C.

As with most K⁺ channels (Rudy, 1988), Rb⁺ had the highest permeability ratio of 0.53, NH₄⁺ was much less permeant with a ratio of 0.08, Cs⁺ was slightly more permeant than in other K⁺ channels with a ratio of 0.04, while Na⁺ was virtually impermeant ⁠. From these experiments, we determined that Cs⁺, a common K⁺ substitute, should not be used on squid ORNs because the small Cs⁺ current through open K⁺ channels may be difficult to distinguish from a small cation receptor current.

In addition to using ion substitution to study receptor currents in isolation, it is often essential to block voltage-gated channels pharmacologically. While some types of K⁺ channels show distinct pharmacological profiles, there are often exceptions to the following general classification: A-type inactivating K⁺ currents (I_A) (Connor and Stevens, 1971) can be blocked by 4-aminopyridine (4-AP); delayed rectifier K⁺ currents (I_K) are sensitive to tetraethylammonium (TEA⁺) (Hille, 1992); the large-conductance (BK) Ca²⁺-activated K⁺ currents (I_K(Ca)) are blocked by charybdotoxin (CTX) (Miller et al. 1985), while the small-conductance I_K(Ca) (SK) channels are blocked by apamin (Blatz and Magleby, 1986). Cd²⁺ blocks many types of voltage-gated Ca²⁺ channels and, therefore, indirectly blocks I_K(Ca). We found that TEA⁺ and CTX blocked I_K and I_K(Ca), respectively, whereas 4-AP mainly blocked I_A, but also blocked I_K. Apamin had no effect. No single agent blocked all outward currents.

In Fig. 3A, the total outward current at a test potential of +40 mV is shown in trace a. Both inactivating and sustained portions of the K⁺ current are evident. The sustained I_K in the presence of 5 mmol l⁻¹ 4-AP is shown in trace b, and the 4-AP-sensitive current, I_A, obtained by subtracting currents in the absence and presence of 4-AP, is shown in trace c. The mean 4-AP block of the peak current at +40 mV was 22±5 % (N=8) and ranged from 5 to 41 %. The cell-to-cell variability in the proportion of I_A in the total K⁺ current is reflected in this broad range of percentage block by 4-AP (see also Fig. 10C). To determine whether the block by 4-AP was dependent on voltage or current direction, we used symmetrical 200 mmol l⁻¹ K⁺ solutions and measured I–V relationships over the voltage range from −60 mV to +60 mV (Fig. 3B). The block by 4-AP was not strongly dependent on voltage or current direction. We also found that 5 mmol l⁻¹ 4-AP was not completely selective for the inactivating current but also blocked a portion of the sustained current (measured at the end of the voltage step). Block of both currents by 4-AP was reversible, as shown in a different cell in Fig. 3C. In most cells, the 4-AP-sensitive currents recovered quite well even after relatively long (10 min) exposure times.

The sustained, delayed rectifier-type K⁺ currents were reversibly blocked by external application of TEA⁺ (Fig. 4A,C) but I_A persisted in the presence of 5 mmol l⁻¹ TEA⁺ (Fig. 4A, trace b). The sustained TEA⁺-sensitive current I_K obtained by subtraction is shown in Fig. 4, trace c. The I–V relationships for I_K shows that the block by TEA⁺ occurs over the voltage range from −60 to +60 mV (Fig. 4B). As with I_A, the TEA⁺-sensitive component of the total current (I_K) varied between cells, with a range of 24–80 % and an average value of 54±6 % (N=9).

Application of 200 nmol l⁻¹ CTX blocked I_K(Ca) (Fig. 5A) over a voltage range of −60 mV to +110 mV (Fig. 5B). The K⁺ currents that were not sensitive to CTX are shown in Fig. 5B, trace b, while the CTX-sensitive current is shown in trace c. These experiments were performed in normal ASW and revealed that, as the voltage approached the Ca²⁺ reversal potential, the CTX-sensitive current decreased as expected for I_K(Ca). On average, 200 nmol l⁻¹ CTX blocked 46±12 % (N=5) of the total sustained current with a range of 28–73 %. Fig. 5C shows that even with a relatively long washout time (approximately 8 min), the block by CTX was not completely reversible. Apamin, which blocks SK in other cells (Blatz and Magleby, 1986), had no effect on the K⁺ currents in squid ORNs (N=6; data not shown), suggesting a uniform population of large-conductance channels.

In addition to CTX, 10 mmol l⁻¹ Cd²⁺ was used to block voltage-gated Ca²⁺ currents which should have the effect of reducing I_K(Ca). Trace a in Fig. 6A shows the total current from a cell that did not have an obvious I_A. In the presence of 10 mmol l⁻¹ Cd²⁺, the block of I_K(Ca) revealed a pronounced inactivating current (Fig. 6A, trace b). The subtracted Cd²⁺-sensitive current in trace c shows both a small transient outward peak and a slower sustained outward current. The transient peak may be due to a direct slowing effect of Cd²⁺ on K⁺ channel activation (Gilly and Armstrong, 1982). The Cd²⁺-sensitive sustained current consists of indirect inhibition of I_K(Ca) by blocking Ca²⁺ channels and direct K⁺ channel block by Cd²⁺, as reported in rat ORN K⁺ channels (Lynch and Barry, 1991a). The I–V relationships over the voltage range from −60 mV to +100 mV show that the Cd²⁺-insensitive I–V relationship is linear (Fig. 6B, trace b), while the Cd²⁺-sensitive I–V relationship reaches a plateau (Fig. 6B, trace c) near the Ca²⁺ equilibrium potential, suggesting a mixture of I_K(Ca) and I_K. Fig. 6C shows that Cd²⁺ block was reversible in this cell, although some cells showed less complete recovery. The average block of the sustained current by 10 mmol l⁻¹ Cd²⁺ was 40±5 % (N=9). The range in the percentage of current blocked by Cd²⁺ (26–69 %) was similar to the that occurring in 200 nmol l⁻¹ CTX (44–73 %), although the additional effects of Cd²⁺ on the activation kinetics and I_K make it less useful for isolating I_K(Ca).

An experiment in which we examined the additive effects of TEA⁺, 4-AP and Cd²⁺ is shown in Fig. 7. Application of 5 mmol l⁻¹ TEA⁺ dramatically reduced the total current and revealed the peak of the transient I_A (Fig. 7, trace b of inset). Addition of 5 mmol l⁻¹ 4-AP in the presence of 5 mmol l⁻¹ TEA⁺ blocked the transient portion of the current (Fig. 7, trace c of inset). Addition of 10 mmol l⁻¹ Cd²⁺ in the presence of both TEA⁺ and 4-AP removed the Ca²⁺-activated portion of the K⁺ current, so that only a small outward current remained (Fig. 7, trace d of inset).

Measurements of the kinetics and voltage-dependence of the voltage-gated K⁺ channels in squid ORNs were made for use in future modelling of the effects of mixtures of hyperpolarizing and depolarizing receptor currents on the amplitude and frequency of action potentials. We took a number of steps to ensure good voltage-clamp control for kinetic experiments. First, we reduced [K⁺]_i to 200 mmol l⁻¹ to reduce the size of K⁺ currents and thereby to reduce series resistance (R_S) errors which become significant with large currents. Second, only cells with uncompensated R_S2 below 12 MΩ were used in kinetic analyses and in every case 80–90 % of R_S2 was electronically compensated to reduce R_S2 to 2 MΩ or below. Third, the cells were cooled to 10 °C to slow channel kinetics. Our kinetic analyses of outward K⁺ currents revealed two distinct current types, a transient I_A current and a delayed rectifier current (I_K). We did not attempt to separate I_K(Ca) from I_K for the kinetic studies because inclusion of Cd²⁺ to block I_K(Ca) affected I_K kinetics (see Fig. 6A) and because we did not have enough CTX to include it routinely in perfused solutions.

Fig. 8A shows a superimposed family of currents in response to 4000 ms voltage steps from −60 to +60 mV in 20 mV increments. The currents have both rapidly inactivating and sustained phases that represent I_A and I_K, respectively. A 400 ms prepulse to either −90 mV (Fig. 8B, trace a) or −40 mV (Fig. 8B, trace b) before each voltage step was used to separate the transient (inactivating) and sustained currents. Subtracting the two current traces resulted in the −40 mV prepulse-sensitive, rapidly inactivating current, I_A (Fig. 8B, trace c), which activates faster than I_K, as can be seen when the current traces from Fig. 8B are scaled to the same peak current in Fig. 8C. In Fig. 8D, the mean time taken to reach half of the peak current (t_1/2) is plotted against the voltage used to elicit the current (N=6). The faster activation kinetics of I_A were observed at voltages ranging from 0 to +60 mV. At +40 mV, the mean value of t_1/2 for I_A (4.3±0.6 ms; Fig. 8D, trace c) was faster than those for I_T (t_1/2=8.7±1.3 ms; Fig. 8D, trace a) and I_K (t_1/2=13.8±1.1 ms; Fig. 8D, trace b).

I–V relationships for the total current (I_T), the prepulse-insensitive I_K and the subtracted prepulse-sensitive I_A show that, in normal ASW, K⁺-selective currents are outwardly rectifying and activate near −40 mV (Fig. 9A). The I–V relationships from six cells were converted to current densities by dividing the peak current at each voltage by the capacitance of the soma (mean value 9.0±0.6 pF, N=26). The current density versus voltage relationships were averaged and converted into conductance density versus voltage relationships by dividing the peak current density at each potential by the driving force (V−E_rev) according to Ohm’s law. The averaged conductance density–voltage (I–G) relationships were fitted with the Boltzmann equation to obtain the maximum conductance density (Gmax), the voltage at which half of the conductance is activated (V_1/2) and the slope of the voltage-dependence (k). The averaged values of each parameter for I_T, I_K and I_A in normal ASW are shown in Table 1. We found that I_A made up approximately 25 % of the total conductance while I_K made up approximately 75 %. I_A activated at slightly more negative potentials than I_K. The values for k were in the normal range for K⁺ channels and are not significantly different from each other (t-test, P>0.05).

Inactivation kinetics of squid ORN K⁺ currents are rather complicated because both I_A and I_K inactivate, but on very different time scales (see Fig. 8A). We generated the steady-state inactivation curve (h_∞) of I_A by applying the voltage protocol shown in the inset of Fig. 10A, and measuring the peak current recorded in response to the test pulse for each of the 400 ms conditioning pulses (Fig. 10A). The averaged normalized h_∞ curve is shown in Fig. 10B. A Boltzmann fit of the averaged data yielded a half-inactivation voltage for I_A of −42±16 mV and a k value of 16±3 (N=5). The I_A portion of the total current was completely inactivated between 0 and +10 mV. The proportion of I_A in squid ORNs varied from cell to cell, resulting in total outward K⁺ currents that showed varying degrees of inactivation. Fig. 10C shows an example of the variability of the contribution of I_A to the total current from two different cells scaled to the same peak. The time constants for inactivation of I_T, I_A and I_K were obtained by fitting a single exponential to the inactivating portion of prepulse subtracted currents, separated as shown in Fig. 8B. The bar graph in Fig. 10D shows that, at +60 mV, inactivation occurred with time constants of 80±10 ms for I_A, 550 ms ±120 ms for I_K and 190±40 ms for I_T (N=5).

The recovery from inactivation of I_A was dependent on both time and voltage. We studied the time-dependence of recovery from inactivation with a two-pulse recovery protocol separated by a hyperpolarizing voltage step that increased in duration from 25 to 350 ms (Fig. 11A, inset). An example of superimposed current responses from the recovery protocol with a hyperpolarizing step to −100 mV is shown in Fig. 11A. The peak currents of the recovery pulse to +60 mV were normalized to the peak current of the test pulse to +60 mV and plotted as the open squares in Fig. 11C. To study the voltage-dependence of recovery from inactivation, we repeated the recovery protocol with the hyperpolarizing voltage steps ranging from −60 to −100 mV, for a duration of 350 ms (Fig. 11B). The peak currents of the recovery pulses following different hyperpolarizing voltages were normalized to the peak current of the test pulse, and the proportion of current recovered was plotted as a function of recovery duration (Fig. 11C). We found that recovery from inactivation was fastest and most complete at more negative potentials. At −80 mV, approximately 35 % of the inactivating current had not recovered after 350 ms. Partial inactivation of I_A at the normal holding potential of −80 mV may account for the discrepancy between the contribution of I_A to I_T measured using 4-AP or conductance density (approximately 25 %), compared with measurements of steady-state inactivation (approximately 40 %).

Because I_A activated and inactivated faster than I_K, we tested whether deactivation of I_A could be distinguished from deactivation of I_K. To measure the deactivation of I_A in isolation, we used a 400 ms prepulse to either −90 mV or −40 mV before each 2 ms activation pulse to +40 mV and obtained the prepulse-sensitive I_A by subtraction (Fig. 12A). To study the deactivation kinetics of I_K, a 400 ms prepulse to −40 mV was followed by a 50 ms activation pulse (Fig. 12B) instead of a 2 ms pulse, because very little I_K activated during the 2 ms activation pulse (data not shown). I_A deactivates faster than I_K at −120 mV as seen by the plots in Fig. 12C, where we scaled the tail currents from Fig. 12A,B to the same peak. We quantified channel deactivation kinetics in response to the 2 and 50 ms activation pulses at repolarization voltages ranging from −130 mV to −90 mV by fitting single exponential functions to each trace and measuring the time constants. The time constants from the 50 ms protocol (filled circles in Fig. 12D) represent deactivation of I_K and were significantly different (paired t-test; P<0.05) from the faster time constants of I_A (filled triangles in Fig. 12D). These data show that the faster I_A deactivation kinetics can be separated from I_K at all tested voltages.

All of the experiments described above were performed at 10 °C to slow down channel kinetics so that accurate measurements could be made. Because experiments on olfactory receptor neurons in other species were performed at various temperatures, we measured the Q₁₀ of the total K⁺ current by steadily increasing the temperature of the bath from 10.0 to 25.0 °C at a rate of 1 °C min⁻¹. Fig. 13A shows a superimposed family of traces obtained by repeatedly stepping the membrane voltage from −80 mV to +60 mV for 200 ms as the bath temperature increased. The size and shape of the total current changed dramatically with the increase in temperature. To quantify these changes, we examined the effects of temperature on activation and inactivation kinetics of I_T, I_A and I_K at +60 mV. Fig. 13B shows I_T activated at 10 and 22 °C with a −90 mV prepulse and +60 mV test pulse, as described for Fig. 8B. The currents have been scaled to the same peak to demonstrate the changes in kinetics with temperature. Fig. 13C shows scaled I_K at the two temperatures obtained by applying a prepulse to −40 mV. Fig. 13D shows the effects of temperature on I_A and was obtained by subtracting the traces in Fig. 13C from those in Fig. 13B and scaling the peaks. The effects of temperature on activation and inactivation kinetics are summarized in Table 2. Increasing temperature increased inactivation kinetics more than activation kinetics. The Q₁₀ values for activation were similar to the Q₁₀ values of 2–4 for ion channel gating described in axons (Hille, 1992). The larger Q₁₀ values for I_T and I_K inactivation may in part reflect the difficulty in fitting the slowly inactivating components of these currents at 10 °C.

Squid ORNs are capable of responding to odorants with depolarizing or hyperpolarizing receptor potentials (Lucero et al. 1992). To understand how receptor potentials are coded in the shape, amplitude and frequency of action potentials, it is essential to understand the biophysical properties of the K⁺ channels involved. In addition, the K⁺ channels themselves may underlie depolarizing receptor potentials (closing K⁺ channels) or hyperpolarizing receptor potentials (opening K⁺ channels). Finally, knowledge of the pharmacology and selectivity of the K⁺ channels allows manipulation of conditions so that receptor currents can be studied in isolation.

We have characterized the outward currents in whole-cell voltage-clamped pyriform olfactory receptor neurons from the squid Lolliguncula brevis and identified at least three types of K⁺ currents: a rapidly inactivating K⁺ current (I_A), a delayed rectifier K⁺ current (I_K) and a Ca²⁺-activated K⁺ current (I_K(Ca)). An inwardly rectifying K⁺ current was not observed. Although K⁺ currents and/or channels have been characterized from the ORNs of a number of vertebrates, including salamanders Salamandra salamandra (Trotier et al. 1993) and Ambystoma tigrinum (Firestein and Werblin, 1987), toad Caudiverbera caudiverbera (Delgado and Labarca, 1993), frog Xenopus laevis (Schild, 1989), catfish Ictalurus punctatus (Miyamoto et al. 1992), zebrafish Danio rerio (Corotto et al. 1996), mouse (Maue and Dionne, 1987) and rat (Lynch and Barry, 1991a,b,c; Trombley and Westbrook, 1991), K⁺ currents have been characterized from only three invertebrate ORNs, namely lobster Panulirus argus (McClintock and Ache, 1989), moth Manduca sexta (Zufall et al. 1991) and a different species of squid, Loligo opalescens (Lucero et al. 1992).

Relative permeability studies were motivated by the desire to find an impermeant monovalent cation substitute for K⁺. By performing bi-ionic experiments, we found that squid ORN K⁺ channels follow the Eisenman selectivity sequence IV of K⁺>Rb⁺>NH₄⁺>Cs⁺>Na⁺ (Eisenman, 1962), which is similar to that for other K⁺ channels. The major difference between the selectivity of squid ORN K⁺ channels and other K⁺ channels was the decreased permeability ratio for Rb⁺. In frog, P_Rb/P_K is 0.91 and in snail neurons it is 0.74 (Rudy, 1988), whereas in squid ORNs, P_Rb/P_K is 0.53. We also found that, although Cs⁺ had a very low permeability ratio of 0.04, the permeability was not zero. This is important because many studies on the olfactory cyclic-nucleotide-gated channel are performed with Cs⁺ replacing K⁺ on the assumption that Cs⁺ is impermeant through K⁺ channels.

Rapidly inactivating K⁺ currents (I_A) have been described in ORNs of most species studied. I_A appears to be absent from neonatal rat (Trombley and Westbrook, 1991), toad (Delgado and Labarca, 1993) and lobster (McClintock and Ache, 1989). Our average conductance density plots revealed that, at a holding potential of −80 mV, I_A made up approximately 25 % of the total current (Fig. 9B). The I_A described in this study was sensitive to 4-AP. The mean block of the total peak current by 5 mmol l⁻¹ 4-AP was 22 %, and a small portion of I_A remained in the presence of 4-AP. In adult rat ORNs, the block of I_A by 5 mmol l⁻¹ 4-AP was also termed incomplete (Lynch and Barry, 1991a). In the present study, 4-AP blocked a portion of the sustained outward current. In zebrafish ORNs, 5 mmol l⁻¹ 4-AP also blocked a small portion of the sustained current, but the effects on I_A were much greater than those observed in the present study (Corotto et al. 1996). In ORNs of the squid Loligo opalescens, 5 mmol l⁻¹ 4-AP blocked 50 % of the total peak current, but the effects on the sustained current were not investigated (Lucero et al. 1992). Therefore, in Lolliguncula brevis ORNs, as in other species, 5 mmol l⁻¹ 4-AP is neither a complete nor a selective blocker of I_A.

On average, the steady-state inactivation curve showed that I_A contributed 40 % to the total current, which is slightly larger than the contribution of I_A as determined by prepulse subtraction or by 4-AP inhibition studies. The partial inactivation of I_A at −80 mV in the two latter measurements, or the inclusion of some inactivating I_K in steady-state experiments, or both, could account for this discrepancy. As in ORNs from other species, I_A activated at approximately −50 mV, which is more positive than I_A in many non-olfactory cells (−65 mV) (Rudy, 1988) and reached half-inactivation at approximately −40 mV. Recovery of I_A from inactivation was very dependent on time and voltage; even with the longest recovery period used (350 ms), recovery from inactivation was incomplete for every recovery voltage positive to −100 mV. The voltage and kinetic properties of I_A described above suggest that, at a resting potential of −60 mV, between 20 and 40 % of I_A will be inactivated. The activation of I_A is fast enough to contribute to repolarization of the membrane during an action potential. I_A could play a role in setting the interspike interval by activating and inactivating during a receptor potential depolarization (Hille, 1992).

Squid ORNs have a delayed rectifier-type I_K that inactivates slowly compared with I_A. External application of 5 mmol l⁻¹ TEA⁺ reversibly blocked 54 % of I_T and had very little effect on I_A, which is similar to the block by TEA⁺ of Loligo opalescens ORNs (Lucero et al. 1992). We did not attempt to determine directly whether TEA⁺ also partially blocked I_K(Ca), although application of TEA⁺+Cd²⁺ led to a greater reduction in the total current than application of TEA⁺ alone (Fig. 7), suggesting that Cd²⁺ blocked an additional component of the current. Both TEA⁺ and 4-AP have been shown to be behaviorally aversive olfactory stimuli to squid (Gilly and Lucero, 1992). In Loligo opalescens, it appears that as little as 20 % block of the total K⁺ current dramatically increases excitability and affects the shape of the action potential (Lucero et al. 1992). It is likely that, in Lolliguncula brevis, 4-AP and TEA⁺ would be detected in a similar fashion.

The third current that we identified was I_K(Ca). We found that 200 nmol l⁻¹ CTX and 10 mmol l⁻¹ Cd²⁺ blocked similar amounts of I_T, although CTX was more specific for I_K(Ca). As with I_A, the amount of I_K(Ca) varied considerably from cell to cell, with the block ranging from 26 to 73 %. I_K(Ca) could play a role in hyperpolarizing the membrane following a burst of action potentials. In addition, squid neurons respond to at least two odorants (dopamine and betaine) with an increase in [Ca²⁺]_i and a hyperpolarization that inhibits the firing of action potentials (Lucero et al. 1992, 1995). Our recent studies indicate that the inhibitory responses to these odorants are mediated by a Ca²⁺-activated Cl⁻ conductance (see below), although this does not rule out the possibility that I_K(Ca) may mediate hyperpolarizing responses to other odorants, as has been shown in the toad Caudiverbera caudiverbera (Morales et al. 1995).

The three current types described above account for the majority of the outward currents in Lolliguncula brevis ORNs. In most cells, replacement of internal K⁺ with tetramethylammonium completely eliminated outward currents (data not shown), while in a subset of cells, a small outward current remained. This outward current could be either a small outward K⁺ current due to incomplete dialysis of the K⁺-free patch pipette solution with the cytosol or a small voltage-dependent Cl⁻ current that may not be present in every cell. Because of the random appearance of this current, we have not attempted to block it with Cl⁻ channel blockers. As has been described in many species, Lolliguncula ORNs do have an odor-activated Cl⁻ conductance (M. T. Lucero and N. Chen, unpublished observations), but it is not voltage-dependent. Future studies will determine whether two Cl⁻ conductances (voltage-dependent and odor-dependent) are present in squid ORNs.

This study is the first to investigate the temperature-dependence of K⁺ channels in ORNs. To maximize recording conditions and to slow channel kinetics, our experiments were conducted at 10 °C. Most studies on other ORNs have been conducted at room temperature (20–21 °C). To facilitate comparison between our study and others, we made recordings on a few cells while increasing or decreasing the bath temperature from 10 to 25 °C. The effects of increasing temperature on I_T were quite striking. We found that the amplitude, the time to half-peak and the time constant of inactivation showed a temperature-dependence with Q₁₀ values ranging from 2 to 6 (see Table 2). This is slightly greater than the range of Q₁₀ values measured from different ion channels of 2–4 (Hille, 1992), suggesting that these channels are quite sensitive to small changes in temperature. The kinetics of K⁺ currents in giant fiber lobe neurons of Lolliguncula brevis also showed strong temperature-dependence, although a Q₁₀ value was not reported (Horrigan et al. 1987).

The squid olfactory organ is unique among invertebrate and many vertebrate olfactory structures in that the organ is exposed to the external milieu without exoskeletal or cartilaginous coverings. This anatomy makes the organ extremely accessible for both in vivo and in vitro studies of olfaction. Our characterization of the selectivity, pharmacology, kinetics and voltage-dependence of K⁺ currents from the squid Lolliguncula brevis will greatly aid future studies on signal transduction of odor responses and information coding in these cells. This work also increases the number of invertebrate species in which K⁺ currents have been characterized and therefore broadens our basis of understanding action potential generation in olfactory receptor neurons.

We would like to thank Jimmy Lucero for preparing and maintaining the cells and Jonathan Danaceau and David Piper for technical assistance. The squids used in this study were provided by the National Resource Center for Cephalopods (Galveston, TX, USA). We thank NPS Pharmaceuticals, Inc. (Salt Lake City, UT, USA) for the generous gift of charybdotoxin. We also thank David Piper and Dr Mike Michel for critical reading of the manuscript and helpful suggestions. This work was supported by NIH NIDCD grant number DC02587-01 and Office of Naval Research grant number N00014-92-J-1488.