Intracellular-Messenger-Mediated Cation Channels in Cultured Olfactory Receptor Neurons

Manduca sexta females attract their conspecific mates through the release of a unique blend of sex pheromones (Starratt et al. 1979; Tumlinson et al. 1989). The males detect these pheromones with specialized olfactory receptor neurons (ORNs) which innervate long, male-specific sensilla trichodea on the male antennae (Sanes and Hildebrand, 1976a,b; Schweitzer et al. 1976; Keil, 1989; Christensen et al. 1989; Kaissling et al. 1989). After stimulation of pheromone-sensitive ORNs, sensillar potentials, which exhibit several different time constants and which show adaptation after a strong pheromone stimulus, can be recorded extracellularly (Schneider, 1962; Schneider and Boeckh, 1962; Zack-Strausfeld, 1979; Kaissling and Thorson, 1980; Kaissling, 1987; Zack-Strausfeld and Kaissling, 1986; Kaissling et al. 1987; Vogt, 1987). Since the small ORNs are tightly covered by supporting cells and lie beneath a thick cuticle (Keil and Steinbrecht, 1987; Keil, 1989), they are relatively inaccessible for intracellular or patch-clamp recordings. Therefore, a primary cell culture system consisting of differentiating, immunocytochemically identifiable ORNs from third-stage pupae of male M. sexta was developed (Stengl and Hildebrand, 1990). Within 2–3 weeks in culture, the ORNs differentiate morphologically and physiologically. They express at least three different potassium channels and a tetrodotoxin (TTX)-blockable sodium channel, but no voltage-gated Ca²⁺ channels (Zufall et al. 1991c). They respond to their species-specific sex pheromone blend by opening nonspecific cation channels (Stengl et al. 1989, 1992a, b). These pheromone-dependent cation channels do not discriminate between Na⁺and K⁺. Their current reverses around 0mV, irrespective of the main external anion, and shows pronounced inward rectification with external Ca²⁺ buffered to 10⁻⁷ mol l⁻¹ (Stengl et al. 1992b). These cation channels were the only channels observed in cultured ORNs to carry inward currents at potentials more negative than the resting potential, with ‘normal’ ionic gradients (see Materials and methods). Previous evidence has suggested that the pheromone-dependent cation channels were second-messenger-dependent (Stengl et al. 1992b).

After exposure to pheromone, cells of insect antennae exhibit slow and long-lasting increases in cyclic GMP concentration (Ziegelberger et al. 1990) and antennal extracts show G-protein-dependent rapid and transient increases in inositol 1,4,5-trisphosphate (InsP₃) concentration (Boekhoff et al. 1990; Breer et al. 1988, 1990). Hence, it was postulated that a G-protein-dependent phosphoinositidase C generating the two second messengers InsP₃ and diacylglycerol (Berridge, 1987; Gilman, 1987; Berridge and Irvine, 1989) was involved in the generation of the rising phase of the receptor potential, while a guanylate cyclase and probably a protein kinase C (Boekhoff and Breer, 1992) might be involved in its declining phase and in the adaptation of the receptor potentials (Stengl et al. 1992a).

This study investigates whether cultured ORNs respond to activation by G-proteins, application of InsP₃, a rise in internal Ca²⁺ concentration or activation by a protein kinase C by opening cation channels that might play an important role in pheromone transduction.

The moths Manduca sexta (Lepidoptera: Sphingidae), reared from eggs on an artificial diet (modified from that of Bell and Joachim, 1976), were kept on a long-day photoperiod regimen (17h:7h light:dark, with lights on at 07:00h) at 25–26°C and 50–60% relative humidity. Pupae were staged as previously described (Sanes and Hildebrand, 1976a; Tolbert et al. 1983). They were usually selected for dissection between 08:00 and 10:00 h and anesthetized by chilling on ice for 10–15min before dissection of the antennal flagellum.

Unless otherwise specified, all culture media were purchased from GIBCO (Grand Island, NY) and all chemicals and biochemicals, from Sigma Chemical Co. (St Louis, MO).

Details of the culture techniques have been reported previously (Stengl and Hildebrand, 1990). Briefly, antennal flagella from male M. sexta pupae (stage 3 of the 18 stages of pupal development) were disrupted by a combination of mechanical and enzymatic treatment. The dissociated cells were plated on concanavalin-A-coated coverslips or uncoated Falcon plastic dishes, in Leibowitz (L15) medium, supplemented with 5% fetal bovine serum (Hyclone) and conditioned medium (supernatant fluid from cultures of a non-neuronal M. sexta cell line, generously provided by Drs J. Hayashi and L. Oland). The cultures were maintained in an incubator for 2–4 weeks at about 20°C and high humidity.

Patch-clamp experiments followed the method described by Hamill et al. (1981). Patch pipettes were made from borosilicate glass capillaries (Clark Electromedical Instruments, Reading, UK) with a Sutter Instruments micropipette puller (model P80/PC). The pipettes were coated with Sylgard (Dow Corning, Midland, MI). The tip resistance was 5–20 MΩ when the electrodes were filled with physiological saline. The cells were viewed at 400× magnification with an Olympus inverted microscope equipped with phase contrast or Hoffmann modulation contrast optics. After formation of a seal between the pipette and the cell membrane, the electrode capacitance was compensated.

Whole-cell currents were measured at room temperature with an Axopatch 1C patch-clamp amplifier (Axon Instruments Co., Burlingame, CA). The currents were acquired on-line with an 80386-based microcomputer (Dell Computer Corp., Austin, TX) using pClamp software (Axon Instruments Co.), which was also used for data analysis. Leakage currents were generally not subtracted since they remained negligible. Only in a few cells (as indicated in the figure legends), where Cs⁺, TTX and Ni²⁺ did not block all the voltage-dependent currents, were leakage currents subtracted. Junction potential drifts (usually around 10pA, as determined by the amplifier) that occurred during some recordings were subtracted with pClamp software. For current–voltage plots, currents were generally measured at least 5ms after the start of the voltage pulse and during the current plateau.

During whole-cell recordings, cultured ORNs were kept in ‘extracellular saline solution’ containing (in mmol l⁻¹): 156 NaCl, 4 KCl, 6 CaCl₂, 5 glucose and 10 Hepes (adjusted to pH7.1 with NaOH). For determination of the ion-specificity of the cation channels, Na⁺was replaced by 156mmol l⁻¹KCl or CsCl; Cl⁻was replaced by acetate or aspartate. During examination of the Ca²⁺ dependence and Ca²⁺ permeability of the cation channels, the cells were bathed in 160mmol l⁻¹ CsCl, 10mmol l⁻¹ Hepes, 5 mmol l⁻¹ glucose, containing 20mmol l⁻¹, 5mmol l⁻¹ or 10⁻⁷ mol l⁻¹ free Ca²⁺(11mmol l⁻¹ EGTA and 1mmol l⁻¹ Ca²⁺). To block Ca²⁺channels, 6mmol l⁻¹ NiCl₂ was added in most experiments. In all experiments, the voltage-dependent Na⁺ channels were blocked with 10⁻⁸mol l⁻¹external TTX and the delayed rectifier K⁺ channels were blocked with 160mmol l⁻¹ internal Cs⁺, while the Ca²⁺-dependent K⁺ channels could not be blocked by Cs⁺ or any other blocker tested.

The ‘intracellular saline solution’ used to fill pipettes was (in mmol l⁻¹): 150 KCl, 5 NaCl, 1 CaCl₂, 11 EGTA, 1.5 MgCl₂, 10 Hepes. For most whole-cell recordings (and all recordings illustrated) KCl and NaCl were replaced by 160 CsCl and MgCl₂ was omitted. To examine the Ca²⁺ dependence of the cation channels, intracellular solutions were used with Ca²⁺ concentrations of 10⁻⁶, 10⁻⁷, 10⁻⁸mol l⁻¹ or of less than 10⁻⁸ moll⁻¹ buffered with BAPTA (Calbiochem, La Jolla CA): 1mmol l⁻¹ BAPTA, 0.9mmol l⁻¹ CaCl₂ (pCa=6); 1mmol l⁻¹ BAPTA, 0.5mmol l⁻¹ CaCl₂ (pCa=7); 1mmol l⁻¹ BAPTA, 0.09mmol l⁻¹ CaCl₂ (pCa=8); or 2mmol l⁻¹ BAPTA, 0.09mmol l⁻¹ CaCl₂ (pCa<8).

‘Normal’ ionic gradients were defined as high [NaCl] externally and high [KCl] internally. The bath solution contained (in mmol l^-1): 156 NaCl, 4 KCl, 6 CaCl₂, 5 glucose and 10 Hepes (adjusted to pH7.1 with NaOH) outside. The solutions in the pipette contained (in mmol l⁻¹): 156 KCl, 5 NaCl, 1 CaCl₂, 11 EGTA (pCa=7), 10 Hepes. For almost all recordings (and for all recordings illustrated), KCl and NaCl were replaced by 160mmol l⁻¹ CsCl.

Before breaking into the whole-cell configuration, the patches were kept at 0mV pipette potential while phorbol esters (TPA, phorbol 12-myristate 13-acetate), or protein kinase C inhibitors such as staurosporine (Boehringer Mannheim, Indianapolis, IN) or H7, were applied to the cell via a glass pipette (tip opening about 10 μm) which was driven by a Picospritzer (General Valve Corp., Fairfield, NJ). All other agents were included in the patch pipette filling solutions, but only in the shaft; the tip of the electrode was filled with about equal parts of the intracellular control solution. Therefore, the concentrations of the agents used are more dilute (to a maximum of 50%) than indicated.

After 2–3 weeks in vitro, ORNs originating from pupal antennae of 3-day-old M. sexta males were examined for the presence of intracellular-messenger-gated cation currents. In whole-cell patch-clamp recordings with 160mmol l⁻¹CsCl, 10⁻⁷mol l⁻¹ CaCl₂ in the patch pipette, and 10⁻⁸mol l⁻¹ tetrodotoxin (TTX) outside (to block all the voltage-dependent Na⁺ and K⁺ channels), no cation currents were elicited (N=21) at potentials between −120mV and 70mV (Fig. 1). This was independent of the principal cation (Na⁺, K⁺ or Cs⁺), the principal anion (Cl⁻, acetate or aspartate) and the external Ca²⁺ concentration. The mean ± ^S.^D. of inward currents elicited at −120mV was −4.8 ±12.2pA (N=21). Extracellular Ni⁺ was added in most recordings to block presumptive Ca²⁺ channels.

Experiments designed to identify intracellular-messenger-dependent cation channels with properties similar to the pheromone-dependent cation channels (Fig. 2, and Stengl et al. 1992b) sought channels that (a) carry inward currents at potentials more negative than the resting potential, with currents less than −25pA at −110mV (exceeding the mean leak currents by about five times) under ‘normal’ ionic gradients; (b) show approximately the same reversal potentials (around 0mV) in various extracellular solutions with different principal cations, irrespective of the main anion outside; (c) show inward rectification with 10⁻⁷mol l⁻¹ Ca²⁺ outside (Stengl et al. 1992b), but linear I/V characteristics with 6mmol l⁻¹external Ca²⁺ (Fig. 2); and (d) may be blockable by tetraethylammonium (TEA⁺) (Stengl et al. 1992b). Criterion b was not always a reliable indication of the presence of intracellular-messenger-dependent cation currents since, in some cells, Ca²⁺-dependent K⁺ currents, which were unaffected by any blocker tested, were superimposed on TEA⁺-blockable cation currents.

For all cells tested, currents at potentials between at least -100mV and +70mV were determined in steps of 10 or 20mV in different extracellular solutions, at various holding potentials. For all recordings shown, 160mmol l⁻¹ CsCl and 10⁻⁸ mol l⁻¹ TTX were present to block the voltage-dependent K⁺ and Na⁺ channels. The ORNs were recorded on-line with a microcomputer, which also generated the voltage protocols (see Fig. 1). This procedure allowed quick determination of the reversal potentials and the rectification properties of the currents in different extracellular salines. It did not allow measurement of the exact time course or the exact maximal amplitude of transient currents, because the data were usually not collected as a continuous time record, but were recorded at different times after obtaining the whole-cell configuration. Leakage currents were generally not subtracted in the following experiments, since they remained negligible in control recordings (Fig. 1).

To test whether activation of G-proteins elicits cation currents with the same properties as the pheromone-dependent cation currents (Fig. 2, and Stengl et al. 1992b), agents that influence G-protein activation were included in the patch pipette during whole-cell recordings.

With 10 μmol l⁻¹ to 1mmol l⁻¹ GTPγS (a non-hydrolysable GTP-analog; Dunlap et al. 1987) in the shaft of the patch pipette (without ATP present), 25% (2/8) of the analyzed ORNs displayed currents greater than −25pA at −110mV. With 10 μmol l⁻¹ to 1mmol l⁻¹ GTP γS+ATP (10 μmol l⁻¹ to 5mmol l⁻¹), the number of responding cells increased, so that 75% (9/12) of all ORNs tested displayed currents greater than −25pA at −110mV (Figs 3, 4A–C), even with 6mmol l⁻¹ NiCl₂ outside (N=3). The reversal potential of the GTP γS and the GTP γS+ATP-dependent currents remained around 0mV if the main cation outside was Na⁺, K⁺ or Cs⁺, regardless of whether Cl⁻ or acetate was the anion (Fig. 3). The currents showed inward rectification with 10⁻⁷ moll⁻¹ Ca²⁺ outside (Fig. 3). GTP γS-dependent cation currents (with or without ATP) were blocked with 20mmol l⁻¹ TEA⁺ in the extracellular solution (N=4). Perfusion with 1mmol l⁻¹ GDPl3S (a non-hydrolysable GDP analog) did not invoke any currents that differed from leakage currents (N=4).

With 100 μmoll⁻¹ to 5mmol l⁻¹ ATP added to the pipette solution, 44% of the ORNs tested displayed nonspecific cation currents (N=22) in the absence of external Ni²⁺(not shown). Thus, ATP alone can elicit cation currents with the same properties as the GTP γS-dependent currents, either directly, or indirectly, in cultured ORNs.

When GTPγS-dependent currents, as well as ATP-dependent currents or GTP γS+ATP-dependent currents, were recorded consecutively in the same cell, a significant dependence of the current amplitude on external [Ca²⁺] became obvious (Fig. 4A–C) when the bath solution was changed from 10⁻⁷ mol l⁻¹ Ca²⁺to 6mmol l⁻¹ Ca²⁺. Irrespective of the monovalent cations present (see Fig. 3), the inward current at a constant negative potential increased transiently and reached a stationary lower current level within less than 2s (Fig. 4B,C). In 10⁻⁷ mol l⁻¹extracellular Ca²⁺ (Figs 3, 4A), the elicited cation currents appeared to be more stable and showed inward rectification. The non-linear I/V curve became linear after several seconds in 6mmol l⁻¹ Ca²⁺ outside (Fig. 4C).

Quantification of the amplitudes of the GTPγS-dependent currents was difficult and dose–response curves could not be obtained reliably since the currents contained a transient component that was not always detected on the non-continuous time record used. A detailed analysis of the kinetics of the currents will be provided elsewhere. The current amplitudes observed at -110mV (measured at least 5ms after the start of the voltage pulse, during the current plateau) ranged from −37 ±17pA (mean ± ^S.^D.) in 6 mmol l⁻¹ Ca²⁺ outside to −86 ±35pA in 10⁻⁷ mol l⁻¹ Ca²⁺, both with 10 μmoll⁻¹ GTP γS (N=6), and from −36 ±18pA in high [Ca²⁺] to −210 ±96pA in low [Ca²⁺], both with 1mmol l⁻¹ GTP γS (N=5).

The possibility that a G-protein-dependent activation of phosphoinositidase C might have caused opening of these GTPγS-dependent cation channels, as suggested by biochemical evidence (Breer et al. 1990), was considered. The cultured ORNs were stimulated with inositol 1,4,5-trisphosphate (InsP₃). If InsP₃ (0.01–100 μmol l ⁻¹ and no ATP) was included in the patch pipette during the whole-cell recordings, cation currents were larger than −25pA at −110mV in 77% (30/39) of all recordings, even in the presence of external Ni²⁺ (Fig. 5). These cation currents also reversed around 0mV. The reversal potential was independent of the principal external cation, and the external anions (Fig. 5). If TEA⁺ was added extracellularly (Fig. 6), no currents were elicited in ORNs with InsP₃ in the patch pipette, regardless of the extracellular solutions used (N=14).

The amplitude and I/V relationship of the InsP₃-dependent currents were also dependent on extracellular [Ca²⁺], in a strikingly similar manner to the GTPγS-dependent currents (Fig. 4). The InsP₃-dependent currents at −110mV ranged from −26 ±12pA (mean ± ^S.^D.) in 6mmol l⁻¹ Ca²⁺ outside to -122 ±119pA in 10⁻⁷ moll⁻¹ Ca²⁺ outside, both with 1 μmol l⁻¹InsP₃ (N=5) and from −21 ±12pA in 6mmol l ⁻¹Ca²⁺ to − 212 ±126pA in 10⁻⁷mol l⁻¹ Ca²⁺, both with 100 μmol l⁻¹ InsP₃ (N=6).

To determine whether InsP₃ opens cation channels via an increase in internal Ca²⁺ concentration, agents known to change the Ca²⁺ levels were included in the patch pipette solution. With Ca²⁺ inside buffered to less than 10⁻⁸ mol l⁻¹ (2mmol l^{− 1} BAPTA+0.09mmol l^{− 1} CaCl₂) during perfusion with InsP₃, only 22% (2/9) of the recorded ORNs displayed cation currents. After increasing the Ca²⁺ concentration in the patch pipette solution to 10⁻⁶ moll⁻¹(BAPTA-buffered, without any addition of other intracellular messengers), cation currents with the characteristics of the InsP₃-and GTPγS-dependent currents appeared in 85% (11/13) of the cells tested (Figs 7–9). The Ca²⁺-dependent currents at -110mV ranged from −148 ±120pA (mean ± ^S.^D.) in 6 mmol l ⁻¹ Ca²⁺ outside ( N=9) to −141 ±136pA in 10⁻⁷mol l⁻¹ Ca²⁺ outside (N=13). Again, these Ca²⁺-dependent cation currents reversed around 0mV, irrespective of the main anion or cation outside (Fig. 7), were TEA⁺-blockable and depended on the extracellular Ca²⁺concentration (Figs 7–9). Switching from low [Ca²⁺] outside to 6 mmol l⁻¹ Ca²⁺ transiently increased the Ca²⁺-activated current (Fig. 8B,C), which then reached a more stable lower current level with a linear I/V relationship (Figs 8C, 9A). The transient current increase was not observed in all recordings (compare Figs 8A–C, 9A).

After switching from 6mmol l ⁻¹ Ca²⁺ outside to 10 ⁻⁷ mol l⁻¹ Ca²⁺, the inward currents increased and became inwardly rectifying (Figs 7, 9A). Switching back to 6mmol l⁻¹ external Ca²⁺, the inward currents increased temporarily to a higher current amplitude (Fig. 8B), before declining again and becoming linear (Figs 8C, 9A). Maximal shifts of about 12mV to more positive values in the reversal potential of the Ca²⁺-dependent cation currents were observed when the extracellular Ca²⁺ concentrations were changed from 6mmol l⁻¹ to 10⁻⁷ mol l⁻¹ (=buffered Ca²⁺) in symmetrical CsCl solutions (Fig. 9B). The mean ± S.D. of the reversal potential shortly after changing the extracellular Ca²⁺ concentration was −7.7 ±4.2mV in high [Ca²⁺] outside (N=12) and −0.3 ±1.5mV in buffered Ca²⁺ outside (N=18).

Finally, the possibility that Ca²⁺ also opens cation channels in cultured ORNs indirectly via activation of a protein kinase C (PKC) was investigated. In 52% of the ORNs tested (N=44), cation currents with a reversal potential around 0mV were elicited when 10pgml⁻¹ phorbol ester (TPA=PKC activator) was applied to the cell via a micropipette before obtaining the whole-cell configuration or with TPA in the patch pipette (Figs 10, 11A,B). In contrast to the InsP₃-, GTP γS-or Ca²⁺-dependent currents, the TPA-evoked currents were not dependent on extracellular [Ca²⁺] (Fig. 11A,B), since they did not show inward rectification in 10⁻⁷ mol l⁻¹ external Ca²⁺. The PKC-dependent cation currents were blocked with 20mmoll⁻¹extracellular TEA⁺ (N=3) (Fig. 10). At −110mV, the PKC-dependent currents ranged from -45 ±16pA in 6mmol l⁻¹ Ca²⁺ (N=6) to −64 ±34pA in buffered Ca²⁺ (N=3).

To test whether the InsP₃-dependent currents were also elicited from a Ca²⁺-dependent phosphorylation via a PKC, PKC inhibitors were applied (Figs 12, 13). With 10μmol l⁻¹ H7 or 2 μmol l⁻¹ staurosporine applied with 10 μmol l⁻¹ InsP₃, 50% (4/8) of the ORNs tested displayed transient cation currents (Fig. 12). The other 50% did not respond. These transient cation currents were elicited only during the first voltage protocol (Fig. 12). In comparison, 77% (30/39) of the ORNs tested expressed more-stable currents without the addition of PKC inhibitors (Fig. 13), as did the TPA-dependent currents (Fig. 10).

The development of a primary cell culture system of differentiating, identifiable ORNs from M. sexta antennae has greatly facilitated studies of the physiological properties of olfactory neurons (Stengl and Hildebrand, 1990). Biochemical evidence indicates the involvement of second messengers in pheromone transduction in these olfactory neurons (Ziegelberger et al. 1990; Breer et al. 1990). Furthermore, cultured ORNs respond to pheromonal stimulation by opening of nonspecific cation channels that appear not to be directly gated by the pheromone. Therefore, the possible occurrence of cation channels mediated by an intracellular messenger was sought.

Whole-cell patch-clamp recordings from the soma of 3-week cultured ORNs showed that GTP γS, ATP, InsP₃, 10⁻⁶ mol l⁻¹Ca²⁺ and PKC cause inward currents of at least −25pA at −110mV, irrespective of the principal monovalent cation or anion outside the cell. Previous experiments have shown that ORNs from antennae of male M. sexta respond to their sex pheromone in vitro with the opening of cation channels that are distinguished from other channels by inward currents at potentials more negative than the resting potential. They have a reversal potential around 0mV, irrespective of the principal external cation or anions, promote linear currents in 6mmol l⁻¹external Ca²⁺(Fig. 2) and express inward rectification in 10⁻⁷ mol l⁻¹external Ca²⁺(Stengl et al. 1992b).

The percentage of pheromone-sensitive cells observed in vitro (38%) (Stengl et al. 1992b) correlates well with the percentage of pheromone-sensitive cells in the antenna, which is about 32% (Lee and Strausfeld, 1990). Because more than 70% of the cultured ORNs responded to intracellular messenger application, it is assumed that ORNs sensitive to plant odors as well as those sensitive to pheromone contain cation channels that are mediated by intracellular messengers. Considering the transient nature of at least some of these channels, they may be present in most, or perhaps all, ORNs. But since at least 2s is required to generate the first computer-driven voltage protocol after breaking into the whole-cell configuration, they might have been overlooked. Furthermore, since these agent-dependent currents possibly depend on intracellular organelles and molecules (such as intracellular Ca²⁺stores), fast washout of the intracellular medium could obliterate the intracellular-messenger-dependent currents. Finally, the presence of intracellular Ca²⁺ buffers could decrease the agent-dependent cation currents in some of the cells to below the levels of leakage currents.

Currents elicited by GTP γS, ATP, InsP₃ and 10⁻⁶ moll⁻¹Ca²⁺ had similar properties and could be blocked by reducing the intracellular Ca²⁺ concentration to less than 10⁻⁸ mol l⁻¹ with BAPTA. Therefore, it is likely that InsP₃, GTP γS and ATP might act via an increase in internal Ca²⁺ concentration, possibly via activation of a G-protein-dependent phosphoinositidase C (Berridge, 1987; Ferris and Snyder, 1992; Gilman, 1987; Stengl et al. 1992a). This has been suggested by biochemical experiments with antennal extracts (Boeckhoff et al. 1990; Breer et al. 1990). It is assumed that InsP₃ might produce a rise in internal Ca²⁺ concentration by promoting Ca²⁺ influx from outside (Restrepo et al. 1990) and/or via release from internal stores, as has been reported in many other systems (Berridge and Irvine, 1989). A release of Ca²⁺ from internal stores in the soma (Berridge and Irvine, 1989) seems likely, since cation channels opened after perfusion with InsP₃ in the presence of external Ca²⁺ channel blockers.

Direct activation of at least a subpopulation of the intracellular-messenger-dependent cation channels by a rise in internal [Ca²⁺] seems probable, since 10⁻⁶ mol l⁻¹ Ca²⁺ elicits cation currents, even in the presence of PKC blockers. Similarly, the ‘spontaneous’ opening of cation channels after patch excision (McClintock and Ache, 1990; Stengl et al. 1992b) in solutions containing a high Ca²⁺ concentration could be explained by Ca²⁺-dependent activation of cation channels. The theory of cation channels directly dependent on [Ca²⁺] is supported by in situ recordings from extruded dendrites of ORNs of the moth Antheraea polyphemus (Zufall et al. 1991b). Calcium-dependent cation channels have also been reported in vertebrate ORNs (Schild and Bischofberger, 1991) and in cultured lobster ORNs sustained inward currents were reported after InsP₃ application (Fadool et al. 1991).

The intracellular-messenger-dependent cation channels of cultured ORNs appear to be permeable to Ca²⁺, since a switch to extracellular solutions containing a high (6–20mmol l⁻¹) Ca²⁺ concentration moved the reversal potentials of the cation currents to more positive values and transiently increased the amplitude of the currents. It is assumed that the influx of extracellular Ca²⁺ through the cation channels causes activation of more Ca²⁺-dependent cation channels, which are later closed (in a Ca²⁺-dependent manner) via an unknown mechanism. The Ca²⁺-dependent rapid inactivation of cation channels, also found in vertebrate ORNs (Zufall et al. 1991a), could constitute a mechanism for rapid termination of the physiological response to odor stimulation. This rapid inactivation has been postulated for pheromone transduction by Kaissling (1972). The transient opening of channels might also underlie the on-responses of ORNs that can follow odor pulses.

It is likely that the rise in internal [Ca²⁺] also activates Ca²⁺-dependent kinases (Nishizuka, 1984). This assumption is supported by the finding that the addition of kinase blockers decreases the number of cells in which intracellular-messenger-dependent cation currents are observed. Furthermore, cation channels in cultured ORNs were opened via stimulation by a protein kinase C. Since the PKC-dependent currents were more stable and were not dependent on extracellular Ca²⁺ concentration (they remained linear at all Ca²⁺ concentrations tested), they might represent a channel population different from the GTPγS-, ATP-, InsP₃-and Ca²⁺-dependent cation currents. As an alternative hypothesis, the transient, directly Ca²⁺-dependent cation channels might change their Ca²⁺-dependence (activation as well as blockage) after PKC-dependent phosphorylation (Ewald et al. 1985). This assumption is supported by the observation that the addition of PKC blockers appeared to render InsP₃-dependent currents more transient and that all agent-dependent currents could be blocked by TEA⁺. Single-channel experiments will distinguish between the two hypotheses.

Since PKC-dependent and Ca²⁺-dependent cation channels have been observed in excised patches from extruded dendrites of ORNs of the moth A. polyphemus, the cation currents probably play an important role in olfactory transduction (Zufall and Hatt, 1991; Zufall et al. 1991b). Despite the assumption of Zufall et al. (1991b) that Ca²⁺-dependent cation channels occur only in inner dendrites, it appears more likely that both channel types occur in outer dendrites. The inner dendritic segment and the soma of the ORNs are tightly wrapped by an auxiliary cell, forming a compartment separate from the outer dendritic segment, which extends uncovered beyond the cuticle into the hairshaft (Keil and Steinbrecht, 1987). The inner dendritic membranes are, therefore, not easily accessible, and extruded dendritic vesicles probably consist of outer dendritic membranes, even when the antennal shank is completely shaved off (Keil and Steinbrecht, 1987; Zufall and Hatt, 1991; Zufall et al. 1991b). Because the in situ recordings from extruded dendrites of ORNs in A. polyphemus were undertaken with 2 mmol l^-1 Ca²⁺ outside, and because the dendrites were preincubated with the pheromone up to an hour before recording (Zufall and Hatt, 1991), transient ion channels would not be detected. Therefore, whether transiently activated cation channels and directly Ca²⁺-dependent cation channels also occur on outer dendrites has yet to be determined. Since all the channels found so far on dendrites of ORNs are either directly or indirectly Ca²⁺-dependent, it has yet to be shown whether and how internal [Ca²⁺] increases in the outer dendrite. So far, no intracellular Ca²⁺ stores have been identified in the outer dendrite (Keil and Steinbrecht, 1987; Keil, 1989). In recent experiments, a transient Ni²⁺-blockable Ca²⁺ current, followed by cation currents, was observed after intracellular messenger application (Stengl et al. 1991). We are therefore investigating whether pheromone triggers the transient opening of both InsP₃-dependent and PKC-dependent Ca²⁺ channels in cultured ORNs.

This work was made possible by the generous support of Drs J. G. Hildebrand and R. B. Levine (University of Arizona, Tucson, USA), in whose laboratories in Tucson these experiments were carried out. I also thank Dr F. Gruber, Dr I. Kuhlmann and Dr W. Rathmayer for providing space and equipment in Konstanz, where this work was completed. I am very grateful to Drs C. Erxleben, U. Homberg, H.-A. Kolb, R. B. Levine, W. Rathmayer and Ms M. Cahill for greatly improving the manuscript. I thank Karim Fuad for help with two figures, Dr L. Oland for generous help with the cell cultures and Dr Oland and Dr J. Hayashi for the supply of conditioned medium. This research was supported by a grant from the DFG [STE 531] and in part by NIH grant AI-23253 to J. G. Hildebrand.