Skip to main content
NCBI home page
The Journal of Physiology logoLink to The Journal of Physiology
. 1999 Feb 1;514(Pt 3):759–768. doi: 10.1111/j.1469-7793.1999.759ad.x

Nicotinic acetylcholine currents of cultured Kenyon cells from the mushroom bodies of the honey bee Apis mellifera

Frank Goldberg 1, Bernd Grünewald 1, Hendrik Rosenboom 1, Randolf Menzel 1
PMCID: PMC2269090  PMID: 9882748

Abstract

  1. Acetylcholine-induced currents of mushroom body Kenyon cells from the honey bee Apis mellifera were studied using the whole-cell configuration of the patch clamp technique. Pressure application of 1 mM acetylcholine (ACh) induced inward currents with amplitudes between -5 and -500 pA.

  2. The cholinergic agonists ACh and carbamylcholine had almost equal potencies of current activation at concentrations between 0·01 and 1 mM; nicotine was less potent. The muscarinic agonist oxotremorine did not elicit any currents.

  3. Approximately 80% of the ACh-induced current was irreversibly blocked by 1 μM α-bungarotoxin. Atropine (1 mM) did not block the ACh-induced current.

  4. Upon prolonged ACh application the current desensitized with a time course that could be approximated by the sum of two exponentials (τ1 = 276 ± 45 ms (mean ± s.e.m.) for the fast component and τ2 = 2·4 ± 0·7 s for the slow component).

  5. Noise analyses of whole-cell currents yielded elementary conductances of 19·5 ± 2·4 pS for ACh and 23·7 ± 5·0 pS for nicotine. The channel lifetimes, calculated from the frequency spectra, were τo = 1·8 ms for ACh and τo = 2·5 ms for nicotine.

  6. Raising the external calcium concentration from 5 to 50 mM shifted the reversal potential of the ACh-induced current from +4·6 ± 0·9 to +37·3 ± 1·3 mV. The calcium-to-sodium permeability ratio (PCa : PNa) was 6·4.

  7. In high external calcium solution (50 mM) the ACh-induced current rectified in an outward direction at positive membrane potentials.

  8. We conclude that Kenyon cells express nicotinic ACh receptors with functional profiles reminiscent of the vertebrate neuronal nicotinic ACh receptor subtype.

The mushroom bodies (MB) are paired neuropile structures in the insect brain and are assumed to be engaged in higher neuronal functions such as multisensory integration, learning and memory. The MBs of the honey bee brain are involved in olfactory learning (Menzel et al. 1974; Erber et al. 1980). They comprise a highly ordered neuropile, densely packed with 170 000 intrinsic elements, the Kenyon cells. Kenyon cells are the third order interneurons of the olfactory pathway converging in the MBs with other sensory and modulatory pathways that are crucial for olfactory learning (Hammer & Menzel, 1995). The voltage-dependent ionic currents of the Kenyon cells have previously been described (Schäfer et al. 1994). Histochemical studies with antisera against nicotinic acetylcholine receptors (nAChRs) suggest the existence of cholinergic synapses onto Kenyon cells. The input elements are at least in part olfactory interneurons, since they show acetylcholinesterase immunoreactivity (Kreissl & Bicker, 1989). Use-dependent modification of cholinergic synapses onto Kenyon cells, or their modulation via modulatory interneurons, may be one cellular mechanism underlying olfactory learning. Therefore, the present study investigated cholinergic currents of the Kenyon cells. The final goal is to understand the role of the Kenyon cells in the neuronal network that underlies olfactory learning and memory in insects.

Nicotinic AChRs have been studied in various insect species, such as the locusts Locusta migratoria (Hanke & Breer, 1986; Tareilus et al. 1990) and Schistocerca gregaria (Albert & Lingle, 1993), the cockroach Periplaneta americana (Beadle et al. 1989), the fruit fly Drosophila melanogaster (Bertrand et al. 1994), and the house fly Musca domestica (Leech & Sattelle, 1992). However, compared with the wealth of information on the vertebrate nAChRs, little is known about the insect nAChRs. The two types of vertebrate nAChRs (neuronal and muscular) have clearly distinct properties, such as relative calcium ion permeability (Yawo 1989; Zhang & Feltz, 1990; Mathie et al. 1991; Sands & Barish, 1991), inward rectification (Mathie et al. 1990; Zhang & Feltz, 1990; Sands & Barish, 1992; Ifune & Steinbach, 1992), and bursting behaviour (Colquhoun & Sakmann, 1985; Mathie et al. 1991; Sargent, 1993). Our results indicate that the physiological properties of the ACh-induced current of the honey bee Kenyon cells closely resemble those of the vertebrate neuronal nAChR. We present for the first time data of an insect nicotinic acetylcholine current in a homogeneous population of identified cells, the Kenyon cells of the honey bee mushroom body.

METHODS

Animals and cell preparation

Honey bee (Apis mellifera) pupae were collected from the comb between days 4 and 6 of pupal development, which lasts 9 days under natural conditions.

Kenyon cells were dissected and cultured following a modified protocol of Kreissl & Bicker (1992). Brains were removed from the head capsule in Leibovitz L15 medium (Gibco BRL) supplemented with sucrose, glucose, fructose and proline (42.0, 4.0, 2.5 and 3.3 g, respectively, for 1000 ml) to reach a final osmolarity of 500 mosmol l−1 (preparation medium). The glial sheath was removed gently and the mushroom bodies were dissected out of the brains. After incubation (10 min) in a calcium-free saline solution (mM: 130 NaCl, 5 KCl, 10 MgCl2, 25 glucose, 180 sucrose and 10 Hepes; pH 7.2), mushroom bodies were transferred back to preparation medium (4 mushroom bodies per 200 μl) and dissociated by gentle trituration with a 100 μl siliconized Eppendorf pipette. Cells were then plated in aliquots of 50 μl on polylysine- (poly-L-lysine hydrobromide, MW 150 000-300 000; Sigma) coated Falcon plastic dishes and allowed to settle and adhere to the substrate for at least 15 min. Thereafter, the dishes were filled with 2.5 ml of culture medium (13% (v/v) heat-inactivated fetal calf serum (Sigma), 1.3% (v/v) yeast hydrolysate (Sigma), 12.5% (w/v) L-15 powder medium (Gibco BRL), 18.9 mM glucose, 11.6 mM fructose, 3.3 mM proline and 93.5 mM sucrose; adjusted to pH 6.7 with NaOH; 500 mosmol l−1) and were kept at 26°C in an incubator at high humidity. Because the mushroom bodies can be mechanically dissected out of the brain and contain somata of Kenyon cells exclusively, the culture contained only Kenyon cells. Under these conditions the cells had somata diameters of 10-15 μm as in the intact mushroom body. They grew a few short, fine and sparsely ramified processes and were viable for up to 6 weeks in culture. For measurements, cells were used between culture days 3 and 10. Processes of those cells chosen for recordings did not overlap with neighbouring neurites.

Electrophysiological techniques

Whole-cell gigaohm seal recordings were performed at room temperature following the methods described by Hamill et al. (1981). Recordings were made using an Axopatch 1D amplifier (Axon Instruments). Pulse generation, data acquisition and analysis were carried out using a TL-1 interface in conjunction with pCLAMP programs (version 6.01, Axon Instruments) running on an AT-type microcomputer. Currents were low-pass filtered with a four-pole Bessel (-3 dB) filter and sampled at various frequencies, depending on the addressed question. Details are given in the respective sections. Voltages were corrected for liquid junction potential; leakage currents were not subtracted. Series resistances ranged between 5 and 10 MΩ and were compensated at 80%. Cell capacitance estimated from the capacitance compensation settings of the amplifier was 2.7 ± 0.4 pF (mean ±s.d., n = 66 cells). Electrodes were pulled from borosilicate glass capillaries (GB150-8P, Science Products, Hofheim, Germany) with a horizontal puller (P87, Sutter Instruments, Novato, CA, USA) and had tip resistances between 4 and 6 MΩ in standard external solution (see below). For data analysis we used the numeric computation software MatLab (version 4.2) with a signal processing toolbox, both from The Math Works Inc. (Natick, MA, USA) for MS Windows 3.1.

Solutions

The bath was continuously perfused at 2 ml min−1 with a standard external solution that consisted of (mM): 130 NaCl, 6 KCl, 4 MgCl2, 5 CaCl2, 160 sucrose, 25 glucose and 10 Hepes-NaOH; pH 6.7, 500 mosmol l−1, which are the physiological pH and osmolarity of the honey bee. The standard internal solution contained (mM): 115 potassium gluconate, 40 KF, 20 KCl, 4 MgCl2, 5 BAPTA tetrapotassium salt, 3 Na2ATP, 0.1 Na2GTP, 6 gluthatione, 150 sucrose and 10 Hepes-bis-Tris; pH 6.7, 490 mosmol l−1. All chemicals were purchased from Sigma.

Transmitter application

The agonists acetylcholine, nicotine and carbamylcholine (all purchased from Sigma) were applied by pressure ejection, using a four-channel pneumatic pump PPM-2 (List Medical Systems). Four glass capillaries (borosilicate glass, 1.0 mm o.d., 0.58 mm i.d., Clark Electromedical Instruments) were bundled with heat-shrinkable tubing and pulled on a modified Kopf vertical puller yielding tip diameters of 1-3 μm for each channel. Ejection times were 50 ms for ramp protocols and up to 16 s for prolonged application. The pipette tip was positioned at a distance of 50-100 μm upstream from the cell in the flux of saline solution. The agonists were applied over the whole of the cell surface by this process, which had been checked previously by adding a few crystals of the dye amaranth (Sigma) to the saline and monitoring the diffusion of the saline pulse. Since the cell diameter is very small, the agonist solution should reach all areas of the cell at almost the same time.

Noise analysis

All recordings used for noise analysis were performed at a holding potential of -95 mV. Signals were filtered at 500 Hz. Stationary ACh-induced current signals were digitized at 1 kHz for a period of 10 s and analysed as follows. Noise analysis power spectral densities were calculated by a fast Fourier transform (FFT) routine on an IBM-compatible microcomputer using a Radix-2 algorithm with a Hamming window as implemented in the MatLab software package (MatLab version 4.2, The Math Works Inc.). Several individual spectra were averaged; the average of ten spectra of control noise (recorded before agonist application) was subtracted from the averaged spectrum in the presence of agonist. This difference yielded the agonist-induced noise. Double Lorentzians were fitted using a Nelder and Mead simplex algorithm from the same software package. For analysis of the elementary current iel the above mentioned stationary current signals were divided into 125 ms blocks. The mean inward current I and the current variance σ2 were calculated for these blocks using the MatLab software package. Plotting σ2 as a function of I and performing a linear regression analysis (Fig. 2E and F) yields iel as the slope of the regression curve (Anderson & Stevens, 1973). For a thorough discussion of noise analysis of electrophysiological signals see DeFelice (1981).

Figure 2. Noise analysis.

Figure 2

Open in a new tab

A, steady-state whole-cell currents elicited by nicotine. Signal sample rate, 1 kHz; filtered at 500 Hz. B, power spectra of nicotine-induced and background membrane noise. C and D, for calculations of mean channel lifetimes, power spectra were fitted with two Lorentzians. E and F, to calculate the elementary conductances, whole-cell current fluctuations were measured during 10 s intervals (sampled at 2 kHz; filtered at 1 kHz) and the variances determined.

Reversal potential determination

The reversal potential was determined using fast voltage ramps from +85 to -95 mV (1.8 V s−1) and from +55 to -95 mV (1.5 V s−1). Since the pharmacological properties of voltage-activated ionic currents of Kenyon cells from the honey bee have previously been described (Schäfer et al. 1994), most of these currents could be selectively blocked. For measurements using voltage ramps, the saline solution contained 100 nM TTX (Sigma) to block voltage-activated sodium currents and 50 μM CdCl2 to block voltage-activated calcium currents. Adding 20 mM tetraethylammonium chloride (TEA-Cl, Sigma) to the pipette solution blocked calcium-activated potassium currents and reduced the delayed rectifier potassium current. At a holding potential of -65 mV the voltage-activated A-type potassium current was only partially inactivated. Therefore, a preceding ramp from -65 to +85 mV (duration, 100 ms) was applied before the test ramps, which completely inactivated the A-type potassium current. Under these conditions, only the rest of the delayed rectifier-type potassium currents and possible leak currents may have overlaid the ACh-induced current. Recordings were performed using voltage ramps with and without an ACh pulse; the difference between the two was obtained through a point-by-point subtraction, and yielded the voltage relationship of the ACh-induced current. Several of these current-voltage curves (I-V curves) of different cells were averaged. The resulting reversal potential was corrected for liquid junction potentials, which were experimentally determined in the current clamp mode of the patch amplifier. For all calculations of permeability ratios, the concentrations of ions were changed to ionic activities at 25°C as described in Robinson & Stokes (1960). For calculations of the calcium ion activities, the Guggenheim convention was used, which relates the mean activity coefficient for CaCl2 (γ±CaCl2) to the individual activity of the calcium ion (γ+Ca2+) by γ+Ca2+ = (γ±CaCl2)2. Values for the mean activity coefficients for CaCl2 at 25°C were obtained from Table 1 in Butler (1968).

RESULTS

ACh-induced current - general features

In the whole-cell mode of the patch clamp technique, pressure application of 1 mM ACh induced inward currents in 163 out of 178 Kenyon cells (91.5%), with peak amplitudes ranging between -5 and -500 pA (121 ± 47 pA, mean ±s.e.m., n = 163). There was no detectable correlation between the maximal amplitude and accessible experimental parameters such as the developmental stage of the pupae used for cell preparation (pupal days 4-6), the time in cell culture, or cell morphology (e.g. size of soma, and number or size of processes).

A rapid onset of the ACh-induced current with a concomitant increase in membrane noise was followed by a fast desensitization (Fig. 1). The time course of desensitization upon prolonged ACh application could be approximated by the sum of two exponentials with time constants ranging between τ1 = 120 and 370 ms (276 ± 47 ms, mean ±s.e.m., n = 21) for the fast component, and between τ2 = 0.9 and 4.2 s (2.4 ± 0.7 s, n = 21) for the slow component. The time constants for the fast and the slow components varied with the type of cholinergic agonist used. Time constants of the desensitization of the carbamylcholine (CCh)-induced current ranged between τ1 = 300 and 650 ms (434 ± 73 ms) for the fast component, and between τ2 = 2.8 and 5.8 s (4.1 ± 0.9 s, n = 19) for the slow component. The time constants of the nicotine-induced current desensitization measured between τ1 = 390 and 420 ms (401 ± 11.4 ms, n = 5) and τ2 = 1.8 and 2.2 s (2.03 ± 0.16 ms, n = 5).

Figure 1. Membrane currents induced by nicotinic agonists.

Figure 1

Open in a new tab

Responses of an individual Kenyon cell to pressure application of the agonists acetylcholine, nicotine and carbamylcholine (duration, 16 s). Inward currents of different amplitudes and time courses were induced by the same agonist concentration (100 μM). The holding potential was -65 mV, and the potential during agonist applications was -95 mV. A strongly calcium-buffered pipette solution was used (5 mM BAPTA/0 mM Ca2+). Sample frequency was 1 kHz; signals were filtered at 500 Hz.

Run-down of the ACh-induced current could be observed in all cells tested. Run-down was characterized by a progressive, irreversible decrease of the current amplitude after obtaining the whole-cell recording configuration. Within 3 min the ACh-induced current reached a steady state. The relative current amplitude at this point was 64 ± 23% (mean ±s.e.m., n = 16) of the amplitude (I0) immediately after obtaining the whole-cell recording configuration. Neither the time course nor the extent of run-down was affected by short pulses of agonist in 10 s intervals. We tried to minimize run-down by adding substances to the pipette solution that are assumed to be necessary for proper functioning of the underlying channels and which could have been washed out of the cell by dialysis with the pipette solution. These substances were glutathione (6 mM), GTP (0.1 mM), cAMP (0.5 mM), cGMP (0.5 mM) and the catalytic subunit of cAMP-dependent protein kinase, which was purified as described by Altfelder & Müller (1991). None of these substances affected the run-down of the ACh-induced current.

Noise analysis

Recordings during prolonged pressure application of 5 μM ACh and 10 μM nicotine (Fig. 2A) were used for noise analysis to estimate the mean channel lifetime and elementary conductance of the underlying channel. At these agonist concentrations desensitization was slow enough to obtain long recordings with sufficiently stable currents (stationary currents). Spectra were obtained by averaging individual spectra of nicotine- (n = 10) and ACh-induced noise (n = 12); from these, averaged spectra of control noise (n = 10) were subtracted (Fig. 2B). The observed frequency spectrum for nicotine could be approximated by a double Lorentzian function with a pronounced low frequency component and cut-off frequencies of fc1 = 64.0 Hz and fc2 = 1.7 Hz (Fig. 2C). The higher frequency component probably represents single channel openings with a time constant of τo = 2.5 ms. The low frequency component represents bursts of openings with a mean duration of 93.2 ms. For ACh-induced noise, the frequency spectrum was also double Lorentzian with cut-off frequencies of fc1 = 90.6 Hz and fc2 = 12.1 Hz, corresponding to single channel openings of τo = 1.8 ms and bursts with a mean duration of 13.6 ms (Fig. 2D).

The elementary current through individual nAChRs was determined by plotting the variance of the agonist-induced noise as a function of the mean inward current and performing linear regression analysis. The elementary current iel corresponds to the slope of the regression curve (Fig. 2E and F). Averaging the results obtained from measurements of nine individual cells yielded an elementary current of -1.85 ± 0.23 pA when ACh was used as the agonist, and -2.25 ± 0.47 pA for nicotine. These elementary currents corresponded to elementary conductances of γ = 19.5 ± 2.4 pS for ACh and γ = 23.7 ± 5.0 pS for nicotine.

Pharmacology

The potencies of the cholinergic agonists ACh and CCh to induce whole-cell currents were almost identical at the concentrations tested, while nicotine was less effective. The peak amplitudes of agonist-induced currents were measured at agonist concentrations of 0.01, 0.1 and 1 mM. Inward currents were detectable after pressure application of 10 μM agonist solution and increased with increasing concentration (Fig. 3). Application of agonist concentrations higher than 1 mM did not further increase the current amplitude (not shown).

Figure 3. Concentration dependency of agonist-induced currents.

Figure 3

Open in a new tab

The experiments were performed on individual cells for each agonist. Due to limitations within the transmitter application system only three concentrations could be tested for each agonist. The inward current amplitudes of a single cell were normalized to the maximum current measured in that cell. The mean relative currents (±s.e.m.) of several cells induced by ACh (n = 10), CCh (n = 8) and nicotine (n = 8) application are plotted.

Pressure application of oxotremorine (1 mM), a potent muscarinic agonist in invertebrates, did not induce membrane currents (n = 7). Bath application of the muscarinic antagonist atropine (1 mM) did not affect the ACh-induced current (not shown). These results define a nicotinic pharmacology of the ACh receptor. The ACh-induced current of Kenyon cells was insensitive to 50 μM cadmium, 100 nM TTX, 10 mM TEA-Cl and 10 mM N-methyl-D-glucamine (NMG) or 10 mM Tris-Cl in the saline solution. The current was completely blocked by 50 μM quinine or quinidine in the external solution (not shown). This block was readily reversible after washing with standard external solution for 30 s to 1 min. Barium (1 mM), zinc (1 mM) or TEA-Cl (20 mM) in the pipette solution had no measurable effect on the ability of cholinergic agonists to elicit whole-cell currents. Incubation with α-bungarotoxin (α-BTX, 1 μM) for 2-5 min blocked approximately 80% of the ACh-induced current (n = 7). Longer incubation periods did not further decrease the current amplitude. The block by α-BTX was irreversible, since no measurable recovery of the current occurred, even after extended periods (several minutes) of washing with saline solution. The α-BTX effect was clearly distinguishable from run-down.

Relative ion permeabilities and rectification

To estimate the I-V relationship of the ACh-induced current, brief (5 ms) pulses of ACh were applied and the peak amplitude was determined at various membrane potentials between -45 and -125 mV (Fig. 4). The I-V curve in this voltage range was approximately linear, and extrapolation to positive membrane potentials revealed a reversal potential between 0 and +10 mV. Since the nicotinic ACh channels are not voltage sensitive, fast voltage ramps were applied to obtain I-V curves over a broad range of membrane potentials (Figs 5 and 6A). When voltage-sensitive currents were pharmacologically blocked, the I-V curve in standard external solution (with 5 mM Ca2+) showed a reversal potential of +4.6 ± 0.9 mV (n = 11, Fig. 5A). Using a modified Goldmann-Hodgkin-Katz (GHK) equation (Jan & Jan, 1976) we calculated that an external concentration of 5 mM Ca2+ should cause a shift in the reversal potential of +3 mV. Accordingly, the measured reversal potential was corrected and a value of +1.6 mV was used to calculate the potassium-to-sodium permeability ratio, PK : PNa = 0.96. Calcium ions in the external solution were required for a stable seal formation; measurements in calcium-free saline were not possible.

Figure 4. I-V relationship of ACh-induced current measured at various potentials.

Figure 4

Open in a new tab

Pressure applications of short (5 ms) ACh pulses at various command potentials (-45, -65, -85, -105 and -125 mV) elicited inward currents of different peak amplitudes (measured at the time indicated by the arrow). The I-V curve obtained is shown in the inset. Holding potential was -85 mV. Signals were sampled at 4 kHz, filtered at 1 kHz and subsequently filtered with a digital filter (Gauss characteristic) at 100 Hz.

Figure 5. I-V curves of ACh-induced currents at different calcium concentrations.

Figure 5

Open in a new tab

Subtraction of currents elicited during a voltage ramp (+85 to -95 mV) without and with ACh application (duration, 150 ms; interval, 550 ms) yielded ACh-induced currents. Signal sample rate, 2 kHz; filtered at 1 kHz. A, I-V curve in standard external solution. B, I-V curve in high calcium solution. Differences in osmolarity were balanced by adjusting the amount of sucrose in the saline. Inset, example of a current trace recorded during the voltage ramps.

Figure 6. Ion substitution experiments.

Figure 6

Open in a new tab

A, replacing external sodium with equimolar NMG reduced ACh-induced current. B, ACh-induced current in the presence of 130 mM sodium (5 mM calcium) at various membrane potentials (inset, I-V curve). C, the same cell showed reduced ACh-evoked current in sodium-free external solution (130 mM calcium). Holding potential, -85 mV; command potentials, -45, -65, -85, -105 and -125 mV. Fast voltage ramps (+55 to -95 mV) applied as shown in Fig. 5.

The reversal potential and the corresponding potassium-to-sodium permeability ratio indicate a cation-selective channel. Replacement of NaCl with N-methyl-D-glucamine chloride (NMG-Cl) in the external solution strongly reduced the amplitude of the ACh-induced current at membrane potentials between -45 and -125 mV (Fig. 6A). The same result was obtained with Tris-Cl instead of NMG-Cl (not shown). This confirmed that sodium and potassium are the main permeant ions. To determine the calcium-to-sodium permeability ratio, ACh-induced currents were recorded during voltage ramps in sodium-free external solution (130 mM NMG-Cl, 5 mM CaCl2). The resulting I-V curve showed a reversal potential of -27.3 ± 1.1 mV (n = 8, Fig. 6A). Using the modified GHK equation, PCa : PNa = 6.4 was calculated, assuming that the ACh receptor is impermeable to NMG (e.g. Zhang & Feltz, 1990). With the above values for PK : PNa and PCa : PNa the modified GHK equation correctly predicts the measured reversal potentials in standard external solution (5 mM CaCl2) and NMG-substituted (sodium-free) external solution. These findings indicate that calcium ions contribute significantly to the ACh-induced current in Kenyon cells, which was further supported by ion substitution experiments. After replacing sodium with equimolar calcium in the external solution, ACh (1 mM) still induced a significant inward current at membrane potentials between -45 and -125 mV (Fig. 6B and C).

In standard saline solution (5 mM CaCl2), the I-V curve displayed a slight non-linearity (Fig. 5A), which became more pronounced at an external calcium concentration of 50 mM (Fig. 5B). Thus, the ACh-induced current showed inward rectification at positive membrane potentials in high external calcium solution. In addition, the reversal potential was shifted to +37.3 ± 1.3 mV (n = 8).

Modulation by intracellular calcium and magnesium ions

The pipette solution used for most experiments contained 4 mM MgCl2 and 5 mM BAPTA. When magnesium was omitted from the pipette solution, ACh did not elicit any currents in Kenyon cells, while voltage-activated currents could still be detected (n = 7). The onset of this effect was rapid; there was no significant ACh-induced current, even shortly after obtaining the whole-cell recording configuration.

Replacing BAPTA with 100 μM calcium in the pipette solution did not impair the ability of ACh to elicit inward currents. The amplitude and time course of the ACh-induced current in the presence of either 5 mM BAPTA or 100 μM calcium were identical (data not shown).

DISCUSSION

Pharmacology

The cholinergic agonists ACh, CCh and nicotine induce inward currents in Kenyon cells of the honey bee mushroom bodies, which can partially be blocked by α-BTX, but not by atropine. Oxotremorine, a muscarinic agonist in insects, does not induce currents. This pharmacological profile clearly defines a nicotinic ACh receptor. Since only about 80% of the ACh-induced current is α-BTX sensitive, two subpopulations of AChR may exist. A large current may flow through α-BTX-sensitive nAChRs and a smaller current through α-BTX-insensitive nAChRs. Similar findings have been obtained by co-expression of different insect AChR α-subunits and chick neuronal β-subunits in Xenopus oocytes (Bertrand et al. 1994). In insects different subpopulations of AChR with different conductance levels and gating kinetics occur on the same cells (Beadle et al. 1989; Leech & Sattelle, 1992; Albert & Lingle, 1993). These were identified as synaptic and extrasynaptic isoforms of the receptor in locust neurons (Tareilus et al. 1990). It may, therefore, be speculated that honey bee Kenyon cells express a synaptic and an extrasynaptic isoform of the nAChR, which are pharmacologically characterized by their different α-BTX sensitivities.

Current through the honey bee nAChR is completely blocked by 50 μM quinine and quinidine. Quinoline derivatives inhibit vertebrate neuromuscular transmission by blocking the muscular nAChR (Sieb et al. 1996). However, no effects of quinoline derivatives on neuronal nAChRs have been reported in either vertebrates or invertebrates. Zinc ions (1 mM) in the pipette solution did not affect ACh-induced currents. In rat neurons, low external zinc concentrations (3-30 μM) inhibit currents through nAChRs (Nutter & Adams, 1995), which may indicate different pharmacologies of the mammalian and the insect nAChRs.

Agonist-induced noise analysis

Single channel data of the honey bee nAChR as derived from noise analyses of whole-cell currents reveal a mean channel lifetime of τo = 1.8 ms with ACh and τo = 2.5 ms with nicotine as the agonist. These values are in good agreement with data from vertebrates (Mathie et al. 1987; Zhang & Feltz, 1990; Mulle et al. 1992). In Kenyon cells the low frequency component of the agonist-induced current may be caused by bursting behaviour of the channel. For the ACh-induced current the corresponding mean burst time of 13 ms is comparable to those of vertebrate neuronal AChRs. The corresponding burst duration of 93.2 ms for nicotine-induced currents may be due to very long bursts or clusters of bursts.

The elementary conductances of the honey bee nAChRs of 20.5 pS with ACh and 25.0 pS with nicotine are in good agreement with vertebrate neuronal nAChR conductances of 20-60 pS (Ifune & Steinbach, 1990; Mathie et al. 1990; Zhang & Feltz, 1990; Mulle et al. 1992; Sands & Barish, 1992) and with single channel conductances of 19-80 pS of insect nAChRs (Beadle et al. 1989; Tareilus et al. 1990; Leech & Sattelle, 1992). However, direct comparison of data from single channel recordings and noise analysis should be made with care. The elementary conductances of the honey bee nAChRs range at the lower end of the spectrum. This may be due to a calcium block of the channel (Imoto et al. 1986) already at a low external calcium level of 5 mM. This block would become more pronounced at higher calcium (50 mM) concentrations, as suggested by the inward rectification of the current at high external calcium concentrations.

Relative ion permeabilities

The honey bee AChR like other nAChRs appears to be a cation-selective channel with sodium and potassium as the main permeant ions and a potassium-to-sodium permeability ratio of 0.96. Its high calcium permeability is the most striking similarity to the vertebrate neuronal AChRs, which show a far greater calcium permeability (PCa : PNa between 0.7 and 20, Sands & Barish, 1991; Adams & Nutter, 1992; Bertrand et al. 1993) than the endplate channel (PCa : PNa = 0.2, Decker & Dani, 1990). However, the permeability ratio (PCa : PNa = 6.4) of the honey bee nAChR may be slightly compromised by a potential small NMG permeability. Equimolar substitution of Ca2+ for Na+ in the external solution reduced the amplitude of inward current through the honey bee nAChR as described in vertebrate neuronal nAChRs (Buisson et al. 1996).

Inward rectification has been shown for a variety of vertebrate neuronal nAChRs (e.g. Yawo, 1989; Mathie et al. 1990; Zhang & Feltz, 1990; Sands & Barish, 1992). The inward rectification of the ACh-induced current of Kenyon cells does not completely abolish currents at positive membrane potentials. This may be explained by assuming that calcium ions bind inside the channel and hinder the permeation of other ions. As was observed, this effect should be especially pronounced at membrane potentials where the net driving force for (monovalent) cations is low, while at higher driving forces (directed inwardly or outwardly) the channel block by calcium ions may be overcome. However, alternative mechanisms such as allosteric effects caused by direct binding of calcium to the receptor cannot be excluded.

Desensitization and run-down

The biphasic desensitization profile indicates two independent processes with different time constants. These probably comprise a fast intrinsic, agonist-induced desensitization (Changeux et al. 1984) and a slower process that may be caused by activity-dependent phosphorylation of the AChR (Huganir & Greengard, 1990). Repeated agonist application resulted in a progressive decrease of current amplitude over seconds, which may be mediated by calcium passing through nAChRs or voltage-sensitive calcium channels and acting either directly or via calcium-activated enzymes. Desensitization of the ACh-induced current in Kenyon cells is rapid and similar to vertebrate α7 and α8 homomers expressed in Xenopus oocytes, whereas α4β2 and α1β1γδ AChRs show slower desensitization rates (Lindstrom, 1996).

Desensitization is clearly distinct from run-down. While desensitization almost completely, but reversibly, abolishes the agonist-induced current in less than 10 s, maximum run-down is reached much later and is not reversible. Desensitization, but not run-down, depends on agonist application, because without agonist application during the first 3 min after obtaining the whole-cell recording configuration, subsequent agonist application did not cause further run-down. Run-down has been observed by several investigators in various experimental settings (e.g. Clapham & Neher, 1984) and may be caused by washout of intracellular substances through dialysis of the cytoplasm with the pipette solution. During this study, supplementing the pipette solution with several substances previously used to prevent run-down (Clapham & Neher, 1984) did not noticeably reduced run-down in Kenyon cells.

Modulation by internal magnesium and calcium

Magnesium ions were required in the pipette solution to elicit agonist-induced currents in Kenyon cells. This is a novel finding and might be due to a direct modulatory effect of magnesium ions on the channel protein. Alternatively, nAChR function in Kenyon cells may require Mg2+/ATP and lack of magnesium ions in the pipette solution may block channel activation. In vertebrates, magnesium may be omitted from the pipette solution, but it induces inward rectification (e.g. Sands & Barish, 1992).

There is contradictory evidence for the effect of intracellular calcium ions on the ACh-induced current. ACh-induced currents did not differ in Kenyon cells measured with an internal solution that contained either 100 μM unbuffered calcium or 5 mM BAPTA. In contrast, the calcium ionophore A23187 reduced the current amplitude (authors’ unpublished observations). However, since A23187 may induce a change in the internal pH, which could influence the current amplitude, this reduction may be calcium independent. Alternatively, the ACh-induced current may be modulated by changes in intracellular calcium, potentially via a calcium-activated protein kinase.

Conclusions

The nicotinic AChR of honey bee Kenyon cells closely resembles the vertebrate neuronal receptor subtype. Specifically, the α7 homo-oligomeric receptor has similar properties with regard to α-BTX sensitivity, calcium permeability, inward rectification, and desensitization rate. One may speculate that one portion of the ACh-induced current passes through an extrasynaptic form of the honey bee nAChR, while the other portion mediates fast synaptic transmission. The involvement of the cholinergic system during memory formation in the honey bee has previously been demonstrated (Gauthier et al. 1992). Although the functional role of the honey bee nAChR during learning and memory formation is still unclear, it is possible that the cholinergic pathway itself provides a modulatory signal. Thus, activation of a presynaptic nAChR may modulate transmitter release as shown in vertebrates (McGehee et al. 1995; Role & Berg, 1996; Wonnacott, 1997). Alternatively, the cholinergic synaptic input into the mushroom body may be regulated by modulatory neurons, given a dendritic localization of the nAChR. Further experiments on the basis of the data presented are planned to explore the function of the honey bee nAChR during olfactory learning.

Acknowledgments

The authors wish to thank Dr Manfred Lindau for critically reading this manuscript. This work was supported by the Deutsche Forschungsgemeinschaft grant SFB 515/C5.

References

  1. Adams DJ, Nutter TJ. Calcium permeability and modulation of nicotinic acetylcholine receptor-channels in rat parasympathetic neurons. Journal de Physiologie. 1992;86:67–76. doi: 10.1016/s0928-4257(05)80009-9. [DOI] [PubMed] [Google Scholar]
  2. Albert JL, Lingle CJ. Activation of nicotinic acetylcholine receptors on cultured Drosophila and other insect neurones. The Journal of Physiology. 1993;463:605–630. doi: 10.1113/jphysiol.1993.sp019613. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Altfelder K, Müller U. Cyclic nucleotide-dependent protein kinases in the neural tissue of the honey bee, Apismellifera. Insect Biochemistry. 1991;21:487–494. 10.1016/0020-1790(91)90102-K. [Google Scholar]
  4. Anderson CR, Stevens CF. Voltage clamp analysis of acetylcholine produced end-plate current fluctuations at frog neuromuscular junction. The Journal of Physiology. 1973;235:655–691. doi: 10.1113/jphysiol.1973.sp010410. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Beadle DJ, Horseman G, Pichon Y, Amar M, Shimahara T. Acetylcholine-activated ion channels in embryonic cockroach neurones growing in culture. Journal of Experimental Biology. 1989;142:337–355. [Google Scholar]
  6. Bertrand D, Ballivet M, Gomez M, Bertrand S, Phannavong B, Gundelfinger ED. Physiological properties of neuronal nicotinic receptors reconstituted from the vertebrate β2 subunit and Drosophila subunits. European Journal of Neuroscience. 1994;6:869–875. doi: 10.1111/j.1460-9568.1994.tb00997.x. [DOI] [PubMed] [Google Scholar]
  7. Bertrand D, Galzi JL, Devillers-Thiery A, Bertrand S, Changeux JP. Mutations at two distinct sites within the channel domain M2 alter calcium permeability of neuronal alpha 7 nicotinic receptor. Proceedings of the National Academy of Sciences of the USA. 1993;90:6971–6975. doi: 10.1073/pnas.90.15.6971. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Buisson B, Gopalakrishnan M, Arneric SP, Sullivan JP, Bertrand D. Human α4β2 neuronal nicotinic acetylcholine receptor in HEK 293 cells: A patch-clamp study. Journal of Neuroscience. 1996;16:7880–7891. doi: 10.1523/JNEUROSCI.16-24-07880.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Butler JN. The thermodynamic activity of calcium ions in sodium chloride-calcium chloride electrolytes. Biophysical Journal. 1968;8:1426–1433. doi: 10.1016/S0006-3495(68)86564-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Changeux JP, Devillers-Thiery A, Chemouilli P. Acetylcholine receptor: an allosteric protein. Science. 1984;225:1335–1345. doi: 10.1126/science.6382611. [DOI] [PubMed] [Google Scholar]
  11. Clapham DE, Neher E. Substance P reduces acetylcholine-induced currents in isolated bovine chromaffin cells. The Journal of Physiology. 1984;347:255–277. doi: 10.1113/jphysiol.1984.sp015065. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Colquhoun D, Sakmann B. Fast events in single-channel currents activated by acetylcholine and its analogues at the frog muscle end-plate. The Journal of Physiology. 1985;369:501–557. doi: 10.1113/jphysiol.1985.sp015912. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Decker ER, Dani JA. Calcium permeability of the nicotinic acetylcholine receptor: the single-channel calcium influx is significant. Journal of Neuroscience. 1990;10:3413–3420. doi: 10.1523/JNEUROSCI.10-10-03413.1990. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. DeFelice LJ. Introduction to Membrane Noise. New York: Plenum Press; 1981. [Google Scholar]
  15. Erber J, Masuhr T, Menzel R. Localisation of short-term memory in the brain of the bee, Apis mellifera. Physiological Entomology. 1980;5:343–358. [Google Scholar]
  16. Gauthier M, Belzunces LP, Zaoujal A, Colin ME, Richard D. Modulatory effect of learning and memory on honey bee brain acetylcholinesterase activity. Comparative Biochemistry and Physiology. 1992;103:91–95. [Google Scholar]
  17. Hamill OP, Marty A, Neher E, Sakmann B, Sigworth FJ. Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflügers Archiv. 1981;391:85–100. doi: 10.1007/BF00656997. [DOI] [PubMed] [Google Scholar]
  18. Hammer M, Menzel R. Learning and memory in the honeybee. Journal of Neuroscience. 1995;15:1617–1630. doi: 10.1523/JNEUROSCI.15-03-01617.1995. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Hanke W, Breer H. Channel properties of an insect neuronal acetylcholine receptor protein reconstituted in planar lipid bilayers. Nature. 1986;321:171–174. doi: 10.1038/321171a0. [DOI] [PubMed] [Google Scholar]
  20. Huganir RL, Greengard P. Regulation of neurotransmitter receptor desensitization by protein phosphorylation. Neuron. 1990;5:555–567. doi: 10.1016/0896-6273(90)90211-w. [DOI] [PubMed] [Google Scholar]
  21. Ifune CK, Steinbach JH. Rectification of acetylcholine-elicited currents in PC12 pheochromocytoma cells. Proceedings of the National Academy of Sciences of the USA. 1990;87:4794–4798. doi: 10.1073/pnas.87.12.4794. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Ifune CK, Steinbach JH. Inward rectification of acetylcholine-elicited currents in rat phaeochromocytoma cells. The Journal of Physiology. 1992;457:143–165. doi: 10.1113/jphysiol.1992.sp019369. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Imoto K, Methfessel C, Sakmann B, Mishina M, Mori Y, Konno T, Fukuda K, Kurasaki M, Bujo H, Fujita Y. Location of a delta-subunit region determining ion transport through the acetylcholine receptor channel. Nature. 1986;324:670–674. doi: 10.1038/324670a0. [DOI] [PubMed] [Google Scholar]
  24. Jan LY, Jan YN. L-Glutamate as an exicitatory transmitter at the Drosophila larval neuromuscular junction. The Journal of Physiology. 1976;262:215–236. doi: 10.1113/jphysiol.1976.sp011593. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Kreissl S, Bicker G. Histochemistry of acetylcholinesterase and immunocytochemistry of an acetylcholine receptor-like antigen in the brain of the honey bee. Journal of Comparative Neurology. 1989;286:71–84. doi: 10.1002/cne.902860105. [DOI] [PubMed] [Google Scholar]
  26. Kreissl S, Bicker G. Dissociated neurons of the pupal honeybee brain in cell culture. Journal of Neurocytology. 1992;21:545–556. doi: 10.1007/BF01187116. [DOI] [PubMed] [Google Scholar]
  27. Leech CA, Sattelle DB. Multiple conductances of neuronal nicotinic acetylcholine receptors. Neuropharmacology. 1992;31:501–507. doi: 10.1016/0028-3908(92)90090-c. [DOI] [PubMed] [Google Scholar]
  28. Lindstrom J. Neuronal nicotinic acetylcholine receptors. Ion Channels. 1996;4:377–450. doi: 10.1007/978-1-4899-1775-1_10. [DOI] [PubMed] [Google Scholar]
  29. McGehee DS, Heath MJS, Gelber S, Devay P, Role LW. Nicotine enhancement of fast excitatory synaptic transmission in CNS by presynaptic receptors. Science. 1995;269:1692–1696. doi: 10.1126/science.7569895. [DOI] [PubMed] [Google Scholar]
  30. Mathie A, Colquhoun D, Cull-Candy SG. Rectification of currents activated by nicotinic acetylcholine receptors in rat sympathetic ganglion neurones. The Journal of Physiology. 1990;427:625–655. doi: 10.1113/jphysiol.1990.sp018191. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Mathie A, Cull-Candy SG, Colquhoun D. Single-channel and whole-cell currents evoked by acetylcholine in dissociated sympathetic neurons of the rat. Proceedings of the Royal Society B. 1987;232:239–248. doi: 10.1098/rspb.1987.0072. [DOI] [PubMed] [Google Scholar]
  32. Mathie A, Cull-Candy SG, Colquhoun D. Conductance and kinetic properties of single nicotinic acetylcholine receptor channels in rat sympathetic neurones. The Journal of Physiology. 1991;439:717–750. doi: 10.1113/jphysiol.1991.sp018690. [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Menzel R, Erber J, Masuhr T. Learning and memory in the honeybee. In: Barton-Browne L, editor. Experimental Analysis of Insect Behaviour. Berlin: Springer; 1974. pp. 195–217. [Google Scholar]
  34. Mulle C, Lena C, Changeux J-P. Potentiation of nicotinic receptor response by external calcium in rat central neurons. Neuron. 1992;8:937–945. doi: 10.1016/0896-6273(92)90208-u. [DOI] [PubMed] [Google Scholar]
  35. Nutter TJ, Adams DJ. Monovalent and divalent cation permeability and block of neuronal nicotinic receptor channels in rat parasympathetic ganglia. Journal of General Physiology. 1995;105:701–723. doi: 10.1085/jgp.105.6.701. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Robinson RA, Stokes RH. Electrolyte Solutions. London: Butterworth; 1960. [Google Scholar]
  37. Role LW, Berg DK. Nicotinic receptors in the development and modulation of CNS synapses. Neuron. 1996;16:1077–1085. doi: 10.1016/s0896-6273(00)80134-8. [DOI] [PubMed] [Google Scholar]
  38. Sands SB, Barish ME. Calcium permeability of neuronal nicotinic acetylcholine receptor channels in PC12 cells. Brain Research. 1991;560:38–42. doi: 10.1016/0006-8993(91)91211-i. [DOI] [PubMed] [Google Scholar]
  39. Sands SB, Barish ME. Neuronal nicotinic acetylcholine receptor currents in phaeochromocytoma (PC12) cells: dual mechanisms of rectification. The Journal of Physiology. 1992;447:467–487. doi: 10.1113/jphysiol.1992.sp019012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Sargent PB. The diversity of neuronal nicotinic acetylcholine receptors. Annual Review of Neuroscience. 1993;16:403–443. doi: 10.1146/annurev.ne.16.030193.002155. [DOI] [PubMed] [Google Scholar]
  41. Schäfer S, Rosenboom H, Menzel R. Ionic currents of Kenyon cells from the mushroom body of the honeybee. Journal of Neuroscience. 1994;14:4600–4612. doi: 10.1523/JNEUROSCI.14-08-04600.1994. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Sieb JP, Milone M, Engel AG. Effects of the quinoline derivatives quinine, quinidine, and chloroquinine on neuromuscular transmission. Brain Research. 1996;712:179–189. doi: 10.1016/0006-8993(95)01349-0. [DOI] [PubMed] [Google Scholar]
  43. Tareilus E, Hanke W, Breer H. Neuronal acetylcholine receptor channels from insects: A comparative electrophysiological study. Journal of Comparative Physiology. 1990;A 167:521–526. doi: 10.1007/BF00190822. [DOI] [PubMed] [Google Scholar]
  44. Wonnacott S. Presynaptic nicotinic ACh receptors. Trends in Neurosciences. 1997;20:92–98. doi: 10.1016/s0166-2236(96)10073-4. [DOI] [PubMed] [Google Scholar]
  45. Yawo H. Rectification of synaptic and acetylcholine currents in the mouse submandibular ganglion cells. The Journal of Physiology. 1989;417:307–322. doi: 10.1113/jphysiol.1989.sp017803. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Zhang ZW, Feltz P. Nicotinic acetylcholine receptors in porcine hypophyseal intermediate lobe cells. The Journal of Physiology. 1990;422:83–101. doi: 10.1113/jphysiol.1990.sp017974. [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from The Journal of Physiology are provided here courtesy of The Physiological Society

RESOURCES