| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Institute of Zoology and Physiology, University of Cologne, Cologne, Germany
Submitted 27 June 2007; accepted in final form 13 November 2007
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
; Strausfeld and Hildebrand 1999; Wilson and Mainen 2006). One experimental system that has served very successfully as a model to understand olfactory information processing is the first-order olfactory relay or antennal lobe (AL) of insects (see Laurent 1999; Wilson and Mainen 2006). As an important step toward the long-term goal to better understand also the cellular mechanisms that mediate olfactory information processing we characterized the physiological and biophysical properties of voltage-activated Ca2+ currents in olfactory interneurons from the ALs of adult Periplaneta americana.
Calcium plays a critical role in the control of a variety of neuronal processes such as synaptic release, membrane excitability, enzyme activation, and activity-dependent gene activation (Augustine et al. 2003; Berridge 1993). A main source of cytoplasmic Ca2+ that contributes significantly to the dynamics of intracellular Ca2+ signals is the Ca2+ influx through voltage-gated Ca2+ channels (VGCCs). Multiple types of voltage-gated Ca2+ channels, characterized by different functional properties, are usually differentially distributed in functionally specialized subcellular compartments of the neuron and contribute to its whole cell Ca2+ current. Characterized by their physiological and biophysical phenotypes the following voltage-gated Ca2+ channel types can be presently distinguished in vertebrates: low-voltage–activated (LVA) channels (T-type, Cav3.1–Cav3.3) and high-voltage–activated (HVA) channels (L-, N-, P/Q-, R-type, Cav1.1–Cav1.4, Cav2.1–Cav2.3).
A comparison of calcium channel gene sequences indicates that in certain structural aspects insect VGCCs might resemble the vertebrate VGCCs (Littleton and Ganetzky 2000). Despite the similarities of Ca2+ channel sequences in invertebrates and vertebrates, invertebrate channels differ greatly in their pharmacological profile. For instance, one of the characteristics of L-type channels in vertebrates is their sensitivity to 1,4-dihydropyridines (e.g., nifedipine), whereas most invertebrate channels with homologous "L-type–like" sequences lack this feature (for review see Jeziorski et al. 2000; Wicher et al. 2001).
In the insect CNS, VGCCs can be separated electrophysiologically into LVA or mid-LVA (M-LVA) and HVA calcium channels (Grolleau and Lapied 1996; Wicher and Penzlin 1997). It has been demonstrated that the LVA current in dorsal unpaired medican (DUM) neurons of P. americana could be further bisected into two components according to their sensitivity to Ni2+ ions: a transient (tLVA) and a maintained LVA current (mLVA; Grolleau and Lapied 1996). In DUM neurons, LVA currents start to activate at –80 mV, M-LVA at –50 mV, and HVA currents at –40 mV. M-LVA and HVA currents were further characterized by their differential sensitivity to inorganic ions (Ni2+, Cd2+) and peptide toxins (conotoxins and agatoxins; Wicher and Penzlin 1997). Differential sensitivity of HVA currents in embryonic brain neurons from P. americana to peptide toxins suggests two current components resembling the vertebrate P/Q- and R-type channels (Benquet et al. 1999).
Organic Ca2+ channel blockers that selectively block specific Ca2+ channel types in vertebrates also modify Ca2+ currents in insects. Earlier studies have demonstrated that Ca2+ currents in insect neurons are reduced by phenylalkylamines (PAAs; e.g., verapamil; Wicher and Penzlin 1997), benzothiazepines (BZTs; e.g., diltiazem; David and Pitman 1995), 1,4-dihydropyridines (DHPs; e.g., nifedipine; Schäfer et al. 1994), and amiloride (Baines and Bate 1998). Investigations in DUM neurons of P. americana showed that HVA currents are affected by verapamil and diltiazem, but not by nifedipine and amiloride (Wicher and Penzlin 1997). However, nifedipine partially blocked barium currents in embryonic brain neurons of P. americana (Benquet et al. 1999). In motoneurons of P. americana Ca2+ current components could be separated by their sensitivity to nifedipine (Mills and Pitman 1997). Detailed analyses, however, of dose–response relationships and effects of organic channel blockers on biophysical properties of Ca2+ currents in insect neurons are still lacking.
The goal of this study was 1) to characterize voltage-activated Ca2+ currents from olfactory interneurons of the antennal lobe in vitro and in situ and 2) to investigate the effects of some Ca2+ channel blockers that have been shown to be effective in vertebrates. The focus for the latter was on verapamil, diltiazem, and nifedipine, all of which selectively block vertebrate L-type channels and are members of different chemical classes. Such detailed knowledge on these readily available blockers could be helpful to block specific components of the calcium currents to better analyze their physiological function in neuronal information processing.
|
METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
P. americana were reared in crowded colonies at 27°C under a 13:11 h light/dark photoperiod regimen and reared on a diet of dry dog food, oatmeal, and water. All experiments were performed with adult animals of both sexes. Before dissection the animals were anesthetized by CO2 or cooling (4°C) for several minutes. For cell culture they were then adhered in a plastic tube with adhesive tape and the heads were immobilized using dental modeling wax (S-U Modelierwachs, Schuler-Dental, Ulm, Germany) with a low solidification point (57°C). For in situ experiments the animals were placed in a custom-built holder, and the head was immobilized with dental wax. The antennae were placed in small tubes on a plastic ring that was later used to transfer the preparation to the recording chamber.
All chemicals, unless stated otherwise, were obtained from Aplichem (Darmstadt, Germany) or Sigma–Aldrich (Taufkirchen, Germany) with a per-analysis purity grade.
Cell culture
To examine the electrophysiological properties of isolated antennal lobe neurons, cells were dissociated and cultured using modified protocols reported previously (Grolleau and Lapied 1996; Hayashi and Hildebrand 1990; Kirchhof and Mercer 1997). The head capsule of anesthetized animals was opened and the antennal lobes were dissected with fine forceps. Typically, ALs from eight animals were pooled in sterile "culture" saline (kept on ice) containing (in mM): 185 NaCl, 4 KCl, 6 CaCl2, 2 MgCl2, 35 D-glucose, 10 HEPES, and 5% fetal bovine serum (S-10, c.c.pro, Neustadt, Germany), adjusted to pH 7.2 (with NaOH), which resulted in an osmolarity of 420 mOsm. For dissociation the ALs were transferred for 2 min at 37°C into 500 µl Hanks Ca2+- and Mg2+-free buffered salt solution (14170, GIBCO, Invitrogen, Karlsruhe, Germany) containing (in mM): 10 HEPES, 130 sucrose, 8 units ml–1 collagenase (LS004194, Worthington, Lakewood, NJ), and 0.7 units ml–1 dispase (LS02100, Worthington), adjusted to pH 7.2 (with NaOH) and to 450 mOsm (with sucrose). Dissociation of neurons was aided by careful trituration with a fire-polished Pasteur pipette for 3–5 min. Enzyme treatment was terminated by cooling and centrifuging the cells twice through 6 ml of culture medium (4°C, 480 g, 5 min). The culture medium consisted of five parts Schneider's Drosophila medium (21720, GIBCO) and four parts Minimum Essential Medium (21575, GIBCO) to which was added (in mM): 10 HEPES, 15 glucose, 10 fructose, 60 sucrose, and 5% fetal bovine serum, adjusted to pH 7.5 (with NaOH) and 430 mOsm (with sucrose). After centrifugation, the cells were resuspended in a small volume of culture medium (100 µl per dish, eight ALs were plated in five to six dishes), and allowed to settle for 2 h to adhere to the surface of the culture dishes coated with concanavalin A (C-2010, Sigma, 0.7 mg ml–1 dissolved in H2O). The cultures were placed in an incubator at 26°C and then used for electrophysiological experiments on the same day. For recordings the cells were visualized with an inverted microscope (IX71, Olympus, Hamburg, Germany) using a x40 water objective (UAPO x40, 1.15 NA, 0.25 mm WD, Olympus) and phase-contrast optics.
Intact brain preparation
The intact brain preparation was based on an approach described by Kloppenburg et al. (1999a,b), in which the central olfactory network was left intact. Shortly before the experiment, the head capsule of the anesthetized animal was opened by cutting a window between the two compound eyes and the bases of the antennae. The brain with antennal nerves and antennae attached was dissected from the head capsule and pinned with fine wire in a Sylgard-coated (Dow Corning, Midland, MI) recording chamber containing "normal" saline (see following text). To gain better access to the recording site and facilitate the penetration of pharmacological agents into the tissue, the brain was enzyme treated (papain, P4762, Sigma, 0.3 mg ml–1 and L-cysteine, 30090, Fluka/Sigma, 1 mg ml–1 dissolved in "normal" saline) for about 3 min at room temperature before the AL was desheathed using fine forceps. The AL neurons were visualized with a fixed-stage upright microscope (BX51WI, Olympus) using a x40 water-immersion objective (UMPLFL x40, 0.8 NA, 3.3 mm WD, Olympus) and IR-DIC optics (Dodt and Zieglgänsberger 1994).
Whole cell recordings
Whole cell recordings were performed at 24°C following the methods described by Hamill et al. (1981). Electrodes (tip resistance between 3 and 5 M
) were fashioned from borosilicate glass (0.86 mm OD, 1.5 mm ID, GB150-8P, Science Products, Hofheim, Germany) with a temperature-controlled pipette puller (PIP5, HEKA Elektronik, Lambrecht, Germany), and filled with a solution containing (in mM): 190 CsCl, 10 NaCl, 1 CaCl2, 2 MgCl2, 10 HEPES, and 10 EGTA adjusted to pH 7.2 (with NaOH), resulting in an osmolarity of 415 mOsm.
During the experiments, if not stated otherwise, the cells were superfused constantly with saline solution containing (in mM): 185 NaCl, 4 KCl, 6 CaCl2, 2 MgCl2, 10 HEPES, and 5 glucose. The solution was adjusted to pH 7.2 (with NaOH) and to 430 mOsm (with glucose).
To isolate the Ca2+ currents we used a combination of pharmacological blockers and ion substitution that has been shown to be effective in other insect preparations (Kloppenburg and Hörner 1998; Kloppenburg et al. 1999b; Schäfer et al. 1994). Transient voltage-gated sodium currents were blocked by tetrodotoxin (TTX, 10–7 to 10–4 M, T-550, Alomone, Jerusalem, Israel). 4-Aminopyridine (4-AP, 4 x 10–3 M, A78403
[GenBank]
, Sigma) was used to block transient K+ currents (IA; nomenclature adapted from Connor and Stevens 1971) and tetraethylammonium (TEA, 30 x 10–3 M, T2265, Sigma) blocked sustained K+ currents [IK(V)] as well as Ca2+-activated K+ currents [IK(Ca)]. In addition the pipette solution did not contain potassium.
Whole cell voltage-clamp recordings were made with an EPC9 patch-clamp amplifier (HEKA Elektronik) that was controlled by the program Pulse (version 8.63, HEKA Elektronik) running under Windows. The electrophysiological data were sampled at intervals of 100 µs (10 kHz), except the 5-ms tail current measurements were sampled at 20 kHz. The recordings were low-pass filtered at 2 kHz with a four-pole Bessel filter. Compensation of the offset potential and capacitance were performed using the "automatic mode" of the EPC9 amplifier. The liquid junction potential between intracellular and extracellular solution (see Neher 1992) of 4.8 mV [calculated with Patcher's PowerTools plug-in from http://www.mpibpc.gwdg.de/abteilungen/140/software/index.html for Igor Pro (WaveMetrics, Portland, OR)] was also compensated. To remove uncompensated leakage and capacitive currents, a p/6 protocol was used (see Armstrong and Bezanilla 1974). Voltage errors due to series resistance (RS) were minimized using the RS compensation of the EPC9. RS was compensated between 30 and 70% with a time constant (
) of 2 µs. Stimulus protocols used for each set of experiments are provided in the RESULTS.
Organic Ca2+ channel modulators
The following organic Ca2+ channel modulators that modify L-type and T-type VGCCs in vertebrate preparations were used in this study: (±)-verapamil (V4629, Sigma), (+)-cis-diltiazem (D2521, Sigma), nifedipine (N7634, Sigma), amiloride (A7410, Sigma), and (±)-BAY K 8644 (B-350, Alomone). (±)-Verapamil and (+)-cis-diltiazem were applied in concentrations ranging from 10–3 to 10–6 M. Nifedipine was dissolved in dimethyl sulfoxide (DMSO; D8418, Sigma) and then added to the saline for final concentrations from 10–4 to 10–6 M. BAY K 8644 was tested at a concentration of 10–4 M. First, it was dissolved in DMSO and then stored in aliquots at –20°C. The aliquot was added to the saline shortly before the experiments were conducted. The DMSO concentration of both the nifedipine- and the BAY K 8644–containing salines was 0.5%. At this concentration it had no obvious effect on ICa and was also added to the "control", although it increased the osmolarity by about 50 mOsm. Due to its photosensitivity (see product sheets from Sigma and Alomone, respectively), all experiments with nifedipine and BAY K 8644 were performed in the dark. Amiloride was dissolved in gently heated saline and used at a concentration of 10–3 M. Drug-containing as well as control saline were bath applied at a rate of 4–7 ml min–1.
Data analysis
The data from the dose–response experiments were fit with a hill equation of the form
 | (1) |
 | (2) |
cpatton/maxc.html; Patton et al. 2004). For analysis of electrophysiological data we used the software Pulse (version 8.63, HEKA Elektronik), Igor Pro 4 (WaveMetrics, including the Patcher's PowerTools plug-ins), and Sigma Stat (Systat Software, Erkrath, Germany). All calculated values are expressed as means ± SD. Significance of differences between mean values were evaluated with Mann–Whitney rank-sum tests or paired t-tests. Significance was accepted at P 
0.05. |
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
500) and the projection neurons (150; numbers from Boeckh and Ernst 1987). It can be expected that the different physiological and computational tasks of these neurons are reflected in the biophysical properties, which might cause a certain variability of the pooled data. If not stated otherwise, the membrane potential was clamped at –60 mV, which is in the range of the normal resting potential of these neurons in situ (–57.2 ± 9.2 mV; n = 28). Typically, depolarizing voltage steps evoked a combination of inward and outward currents in AL neurons. When voltage-gated Na+, K+, and H currents were reduced by ion substitution, pharmacological reagents, and appropriate voltage protocols, we recorded inward currents that had the characteristics of typical Ca2+ currents. Our recording conditions were designed to drastically minimize, or even abolish, outward currents through K+ channels and in most, but not all, recordings we did not observe outward currents. For recordings in which residual outward currents at strong depolarizations were detected we could not rule out that the K+ currents were not completely blocked and that there was some residual K+ in the neurons. However, we think it is more likely that the observed outward currents were carried by Cs+ through Ca2+ channels, which are not completely impermeable to Cs+ (Hess et al. 1986). This scenario was supported by our observation that no residual outward currents could be detected when ICa was blocked by Cd2+ (data not shown). To analyze ICa, outward currents were minimized by substituting K+ in the pipette solution with Cs+ and adding 3 x 10–2 M TEA and 4 x 10–3 M 4-AP to the extracellular solution. The remaining inward current consisted of a transient very fast activating/inactivating component and a more slowly inactivating component (Fig. 1 A), that did not completely inactivate even after long (>1 s) depolarizations (Fig. 1B). By using pharmacological blockers and ion substitution, the main charge carrier of both components could be separated and identified. The fast transient component was a TTX-sensitive sodium current (Fig. 1A), whereas the other components of the inward current were identified as voltage-activated Ca2+ conductances (see following text). To measure ICa, the neurons were superfused with saline containing 10–7 M TTX, 4 x 10–3 M 4-AP, and 3 x 10–2 M TEA. In the pipette solution K+ was replaced with Cs+.
|
Experiments with varying extracellular Ca2+ concentrations confirmed that Ca2+ was the charge carrier of the investigated inward current. Low extracellular Ca2+ concentrations acted quickly to reduce the inward current reversibly (n = 5; Fig. 2 A). This reduction of ICa was concentration dependent. When EGTA containing Ca2+-free extracellular solution was applied, the inward current was completely abolished (data not shown).
|
). Simultaneously the current–voltage (I/V) relation shifted by about 10 mV (n = 4) to more hyperpolarized membrane potentials (Fig. 2D), which might result from changes in the surface charge of the cell membrane (Fedulova et al. 1985). The resulting voltage-dependent differences between ICa and IBa are demonstrated by steps to –20 and –5 mV in Fig. 2, C1 and C2, respectively. Furthermore, the amount of inactivation during a depolarizing voltage pulse was reduced, which became especially obvious during long-lasting depolarizations (1 s; –5 mV; Fig. 2E). These results suggest that inactivation during a voltage step is, at least in part, a Ca2+-dependent mechanism. ICa in vitro
The characteristics of ICa are shown in Fig. 3. The I/V relationship of the peak currents was determined by increasing voltage steps (50 ms, 5 mV) between –60 and 40 mV from a holding potential of –60 mV (Fig. 3A). The voltage dependence of activation of ICa was determined from tail currents that were evoked by 5-ms voltage steps from –80 mV holding potential to 40 mV in 10-mV increments (Fig. 3B). The I/V relations were fit to a first-order Boltzmann equation (Eq. 2; Fig. 3, G and I). Steady-state inactivation of ICa was measured from a holding potential of –60 mV. Prepulses (500 ms) were delivered in 5-mV increments from –95 to –5 mV, followed by a 50-ms test pulse to –5 mV, and the peak currents were determined (Fig. 3C). The I/V relations were fit to a first-order Boltzmann equation (Eq. 2; Fig. 3, H and I).
|
The activation and inactivation kinetics during a voltage step are voltage dependent (Fig. 3, A and B); the time to peak current and the time constant for the decay during a voltage pulse decreased when voltage steps of increasing amplitude were applied. These parameters, however, were not analyzed quantitatively. The tail currents that are independent of the changing driving force during the series of voltage pulses had a maximum amplitude of 1.9 ± 0.4 nA (n = 21). This corresponds to a mean maximal conductance (Gmax) of 12.6 ± 2.7 nS and a mean current density of 63.6 ± 13.8 pA/pF. The I/V relation of the tail currents was fit by a first-order Boltzmann equation with a mean voltage for half-maximal activation (V0.5act) of –17.8 ± 3.3 mV (s = 6.0 ± 2.2; n = 21; Fig. 3, G and I).
Steady-state inactivation started in vitro at prepulse potentials around –60 mV and increased with the amplitude of the depolarizing prepulse (Fig. 3, C, H, and I). The I/V relationship was fit with a first-order Boltzmann equation (Eq. 2) with a voltage for half-maximal inactivation (V0.5inact) of –24.2 ± 3.7 mV (s = 8.7 ± 2; n = 10; Fig. 3, H and I).
ICa in situ
ICa recorded in situ (Fig. 4) showed characteristics similar to those in vitro. However, the I/V relationships had a larger variability and the means were shifted significantly to more depolarized membrane potentials. The mean voltage for activation of the maximal peak current was shifted by 14.4 mV (P < 0.001). The voltage for half-maximal tail-current activation (V0.5act) was shifted by 7.3 mV (P < 0.001). In situ, ICa started to activate at command potentials more depolarized than –40 mV (Fig. 4D). The mean peak currents reached a maximum amplitude (Imax) of 1.2 ± 0.4 nA (n = 22; Fig. 4D) at 8 ± 7.8 mV (Fig. 4E). Based on a mean whole cell capacitance of 28.8 ± 7.8 pF, this corresponds to a mean current density of 42.6 ± 14.3 pA/pF (Fig. 4F). The tail currents had a mean maximum of 1.8 ± 0.4 nA and a mean voltage for half-maximal activation (V0.5act) of –10.5 ± 6 mV (s = 7.5 ± 1.8; n = 13; Fig. 4, G and I). In situ steady-state inactivation started at prepulse potentials around –60 mV and had a mean voltage for half-maximal inactivation (V0.5inact) of –19.9 ± 6.7 mV (s = 8.7 ± 1.9; n = 8; Fig. 4, H and I).
|
In a series of in vitro experiments we analyzed the effect of several inorganic ions and organic substances on ICa, which block or enhance voltage-gated calcium currents in vertebrate preparations. ICa was blocked by the divalent cations Cd2+, Ni2+, and Co2+. We also found that verapamil, diltiazem, and nifedipine, which all belong to different chemical classes (phenylalkalamine, benzothiazepine, and 1,4-dihydropyridine, respectively) and are known to selectively block vertebrate L-type channels, differentially modify ICa. Amiloride (10–3 M), a T-type channel blocker, and (±)-BAY K 8644 (10–4 M), a 1,4-dihydropyridine and L-type channel agonist, had no effect (data not shown).
INORGANIC IONS. The divalent cations Cd2+, Co2+, and Ni2+ blocked ICa in a dose-dependent way. Figure 5, A–C shows single example experiments for each of the blockers and Fig. 5D shows the dose-inhibition curves for all three cations. ICa was elicited by a 50-ms voltage pulse to –5 mV from a holding potential of –60 mV. The dose-inhibition curves were well fit with a Hill equation (Eq. 1). The most effective blocker was Cd2+ with a half-inactivating concentration (IC50) of 10–5 M (Hill coefficient nH = 0.87) followed by Ni2+ (IC50 = 3.13 x 10–4 M; nH = 1.01) and Co2+ (IC50 = 1.06 x 10–3 M; nH = 1.04).
|
VERAPAMIL. The phenylalkalamine verapamil was tested at concentrations ranging from 10–6 to 10–3 M. Example experiments demonstrating the verapamil effect are shown in Fig. 6 A and the dose–response curve is given in Fig. 6A2. An obvious effect of verapamil started at 10–4 M and a nearly complete block was achieved at 10–3 M (Fig. 6). The effects reached a steady state within 3–5 min. The dose-inhibition curve was well fit by a Hill equation (Eq. 1) with a half-maximal block of ICa at 1.5 x 10–4 M (nH = 1.55; Fig. 6A2).
|
The voltage dependence of the peak- and tail-current activation was not modified by verapamil (Fig. 6, C1, C2, and D). However, the voltage dependence for steady-state inactivation of ICa was changed (Fig. 6E): The mean voltage for half-maximal inactivation (V0.5inact) was shifted to hyperpolarized membrane potentials from –27 ± 3.3 mV (s = 7.7 ± 2.2) to –36.4 ± 2.1 mV (s = 6.5 ± 1.5; P = 0.005; n = 9).
DILTIAZEM.
The benzothiazepine diltiazem was tested at concentrations ranging from 10–6 to 10–3 M. Example experiments demonstrating the effect of diltiazem are shown in Fig. 7 A and the dose–response curve is given in Fig. 7A2. The effects reached a steady state within 3–5 min. A clear diltiazem effect started at 10–4 M. The dose-inhibition curve was well fit by a Hill equation (Eq. 1) with a half-maximal block of ICa at a concentration of 2.87 x 10–4 M (nH = 1.06; Fig. 7A2). The ICa block was accompanied by a change in the waveform of ICa (Fig. 7B and inset). During a depolarizing voltage pulse diltiazem increased the rate of inactivation. The mean time constant () of the decay of ICa (from a monoexponential fit) during a voltage pulse to –5 mV was significantly increased from 17.3 ± 2.1 to 28.2 ± 6.5 ms (P = 0.003; n = 6) when the IC50 of diltiazem (3 x 10–5 M) was applied. This effect was even more prominent with higher concentrations of diltiazem (data not shown).
|
NIFEDIPINE. The 1,4-dihydropyridine nifedipine was tested in concentrations from 10–6 to 10–4 M, in which range it was relatively easy to dissolve. Example experiments demonstrating the effect of nifedipine are shown in Fig. 8 A1 and the mean dose–response data are given in Fig. 8A2. An obvious effect of nifedipine on ICa was detectable at 10–5 M (Fig. 8A). The maximal usable concentration of nifedipine (10–4 M) blocked about 33% of ICa (Fig. 8A). The remaining experiments with nifedipine were carried out with a concentration of 10–4 M. At this concentration nifedipine reduced ICa without significantly changing the waveform and the voltage dependence for activation and inactivation (Fig. 8, B–E).
|
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
; Laurent 1999; Strausfeld and Hildebrand 1999; Wilson and Mainen 2006). Together with future research, this work aims to better understand the role of Ca2+ currents in mediating olfactory information processing on both the cellular and the subcellular levels.
ICa recorded in vitro from isolated somata of adult AL olfactory interneurons activated at membrane potentials above approximately –50 mV with a maxiumum current around –5 mV. ICa consisted of both relatively fast activating/inactivating and noninactivating components. In situ the I/V relation for steady-state activation/inactivation was shifted to more depolarized membrane potentials. Interestingly, in a similar study of honeybee antennal motoneurons, differences between in vitro and in situ recordings were not observed (Kloppenburg et al. 1999b). After taking into account the different experimental conditions between in vitro and in situ experiments, however, such a shift of parameters was not unexpected. One reason might be imperfect voltage control across the whole neuron in the intact brain preparations, in which the neuronal arborizations are still intact, and/or differences in voltage dependence of calcium channels that are localized in distal regions of the neurons. Previous studies in other insect and invertebrate preparations have demonstrated neuritic or axonal localizations of calcium channels (Duch and Levine 2002; Haag and Borst 2000; Kloppenburg et al. 2000, 2007). However, given the complex morphology of these neurons it can be assumed that a major part of the measured in situ currents originates from the cell body. Nevertheless, both the in vitro and in situ parameters of ICa are well in the range of Ca2+ currents described in other insect preparations including Drosophila neurons (Byerly and Leung 1988; Saito and Wu 1991), honeybee Kenyon cells (Schäfer et al. 1994), Manduca motor neurons (Hayashi and Levine 1992), embryonic cockroach neurons (Benquet et al. 1999), cockroach DUM neurons (Heidel and Pflüger 2006), locust thoracic neurons (Laurent et al. 1993; Pearson et al. 1993), cricket giant interneurons (Kloppenburg and Hörner 1998), and honeybee antennal motoneurons (Kloppenburg et al. 1999b). The average current density of about 50 pA/pF in vitro was in the same range as that found in vitro for honeybee projection neurons (Grünewald 2003) and cockroach DUM neurons (Heidel and Pflüger 2006).
In this study we presented averaged data from a large number of experiments to characterize the parameter space of ICa in AL interneurons. It is the first stage to characterize ICa of adult AL interneurons in detail and to get pharmacological tools to manipulate ICa. The variability of the data, however, should not be ignored. It might be due to differential expression of Ca2+ channel types in different cell types. This hypothesis can be tested by recordings from neurons that are unequivocally identified by single-cell labeling.
In vertebrates, Ca2+ currents are usually classified into low-voltage–activated (LVA or T-type, with activation starting above approximately –70 mV) and high-voltage–activated (HVA, activation starting above approximately –30 mV) classes. Subtypes such as L-, P/Q-, N-, and R-type are defined by biophysical and pharmacological properties (Ertel et al. 2000; Hille 2001; Triggle 2006). In accordance with previous studies, we found that the pharmacological classification of vertebrate calcium currents is difficult to transfer to ICa of insects (for review see King 2007). The ICa in AL interneurons exhibits some characteristics that are typical for some HVA channel types: Ba2+ is a better charge carrier than Ca2+, inactivation is Ca2+ dependent and relatively slow compared with LVA channels, ICa is more sensitive to Cd2+ than to Ni2+, and ICa is reduced by the L-type blockers verapamil, diltiazem, and nifedipine. However, the activation range of ICa is more hyperpolarized than traditional HVA channels and thus resembles currents with L-type properties that have more mid-voltage–activated ranges (Johnson et al. 2003; Wicher and Penzlin 1997). Despite these "L-type" like characteristics (±)-BAY K 8644 did not modify ICa, indicating that ICa in AL interneurons differs from vertebrate L-type calcium currents pharmacologically. Amiloride, a vertebrate T-type channel blocker, did not affect ICa, whereas amiloride does inhibit calcium currents in different Drosophila preparations [embryonic central neurons (Baines and Bate 1998); larval muscles (Gielow et al. 1995)].
Often the organic Ca2+ channel blockers act more potently on vertebrate cells than on invertebrate neurons and, in previous studies, it has been argued that the ICa blocker concentrations, which are needed to inhibit calcium currents in insect neurons, are too high to achieve any specific effects on calcium channel subtypes (Benquet et al. 2002). Our studies confirmed these concerns. All three blockers—verapamil, diltiazem, and nifedipine—dramatically affected voltage-activated sodium and potassium currents. Similar effects on voltage-activated potassium currents have been described previously (e.g., Caballero et al. 2004; DeCoursey 1995; Trequattrini et al. 1998). Unfortunately, these findings limit the use of the tested blockers for many experimental application. However, some blockers, e.g., diltiazem, reduce a specific component of ICa and thus might be useful for experiments, in which a certain component of ICa is analyzed. To specifically block ICa or its components without affecting other ionic currents different (classes of) substances have to be tested. In this regard spider toxins might provide a more specific tool for ICa characterization (King 2007).
Verapamil
The verapamil-induced block of ICa in AL interneurons is in agreement with studies in cockroach DUM neurons (Wicher and Penzlin 1997), embryonic cockroach brain neurons (Benquet et al. 1999), cockroach motoneurons (Mills and Pitman 1997), and locust thoracic neurons (Pearson et al. 1993). Compared with cockroach DUM and motoneurons the ICa in AL interneurons seems to be less sensitive to verapamil. However, only Benquet et al. (1999) tested different concentrations of verapamil, and this dose-inhibition curve was in the same range as for ICa in this study, although with a slightly lower value for half-maximal inhibition (IC50). Although verapamil did not affect the I/V relationship for activation, it shifted the voltage dependence for the steady-state inactivation of ICa to more hyperpolarized potentials. It was previously reported that this is an effect of drugs that bind to and stabilize the inactivated state of ion channels (Bean et al. 1983; Gomora et al. 2001).
Verapamil, especially at concentrations between 10–4 and 2.5 x 10–4 M, decreased the inactivation rate of ICa in a dose-dependent manner. Considering that in vertebrates various Ca2+ channel subtypes differ in their inactivation kinetics (Budde et al. 2002; Fox et al. 1987), this finding could indicate that ICa in AL interneurons consists of current components with different inactivation kinetics. However, it also has been argued that such an increase in the decay rate could be caused by the blocking mechanism of verapamil ("open channel blocker"; Johnson et al. 1996).
Diltiazem
Similar to the diltiazem-induced block of ICa described here, a reduction of ICa by diltiazem has been demonstrated in cockroach DUM neurons (Wicher and Penzlin 1997) and Drosophila muscles (Gielow et al. 1995). However, dose-dependent effects of diltiazem were not previously investigated in detail.
The current that was not blocked by diltiazem activated and inactivated at more hyperpolarized potentials compared with the control. This finding suggests that a current component activating in the high-voltage range is inhibited by diltiazem, which would be in agreement with previous findings in DUM neurons (Wicher and Penzlin 1997). Compared with our verapamil results, the shift to more negative potentials is less pronounced during the application of diltiazem. Diltiazem, in contrast to verapamil, increased the ICa decay time constant during a depolarizing voltage pulse. Either diltiazem preferentially blocks a current component with fast inactivation kinetics or the drug modifies the gating behavior of a single calcium channel type. Thus diltiazem might be a valuable tool to separate different components of the whole cell ICa of AL interneurons in P. americana.
Nifedipine
Nifedipine reduced the maximal conductance of ICa without affecting the voltage dependence for steady-state activation/inactivation and the decay rate during a voltage step. Thus nifedipine seems not to block a single specific component of ICa, as observed in cockroach motoneurons (Mills and Pitman 1997) and in Drosophila larval muscle (Gielow et al. 1995). However, in this context it is important to consider that most neurons in the present study mainly expressed high-voltage–-activated (HVA) Ca2+ currents, meaning that mostly the effect of nifedipine on HVA Ca2+ channels was examined.
This study provides a detailed biophysical analysis of calcium currents in insect olfactory interneurons. In addition, calcium current pharmacology demonstrated substance-specific effects on ICa for some organic blockers, but also revealed strong nonspecific effects of all tested organic blockers on other voltage-activated currents. The current analysis lays groundwork for our present and future studies in the intact brain to further analyze the basis and role of intracellular Ca2+ dynamics in olfactory information processing, with a focus on cell-type–specific differences.
|
GRANTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
|
ACKNOWLEDGMENTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
|
FOOTNOTES |
|---|
Address for reprint requests and other correspondence: P. Kloppenburg, University of Cologne, Institute of Zoology and Physiology, Weyertal 119, 50931 Cologne, Germany (E-mail: peter.kloppenburg{at}uni-koeln.de)
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
|---|
Augustine GJ, Santamaria F, Tanaka K. Local calcium signaling in neurons. Neuron 40: 331–346, 2003.[CrossRef][Web of Science][Medline]
Baines RA, Bate M. Electrophysiological development of central neurons in the Drosophila embryo. J Neurosci 18: 4673–4683, 1998.
Bean BP, Cohen CJ, Tsien RW. Lidocaine block of cardiac sodium channels. J Gen Physiol 81: 613–642, 1983.
Benquet P, Guen JL, Dayanithi G, Pichon Y, Tiaho F. 
-AgaIVA-sensitive (P/Q-type) and -resistant (R-type) high-voltage-activated Ba2+ currents in embryonic cockroach brain neurons. J Neurophysiol 82: 2284–2293, 1999.
Benquet P, Le Guen J, Pichon Y, Tiaho F. Differential involvement of Ca2+ channels in survival and neurite outgrowth of cultured embryonic cockroach brain neurons. J Neurophysiol 88: 1475–1490, 2002.
Berridge MJ. Cell signalling. A tale of two messengers. Nature 365: 388–389, 1993.[CrossRef][Medline]
Boeckh J, Ernst KD. Contribution of single unit analysis in insects to an understanding of olfactory function. J Comp Physiol A Sens Neural Behav Physiol 161: 549–565, 1987.[CrossRef]
Budde T, Meuth S, Pape HC. Calcium-dependent inactivation of neuronal calcium channels. Nat Rev Neurosci 3: 873–883, 2002.[CrossRef][Web of Science][Medline]
Byerly L, Leung HT. Ionic currents of Drosophila neurons in embryonic cultures. J Neurosci 8: 4379–4393, 1988.[Abstract]
Caballero R, Gomez R, Nunez L, Moreno I, Tamargo J, Delpon E. Diltiazem inhibits hKv1.5 and Kv4.3 currents at therapeutic concentrations. Cardiovasc Res 64: 457–466, 2004.
Connor JA, Stevens CF. Voltage clamp studies of a transient outward membrane current in gastropod neural somata. J Physiol 213: 21–30, 1971.
David JA, Pitman RM. Calcium and potassium currents in the fast coxal depressor motor neuron of the cockroach Periplaneta americana. J Neurophysiol 74: 2043–2050, 1995.
DeCoursey TE. Mechanism of K+ channel block by verapamil and related compounds in rat alveolar epithelial cells. J Gen Physiol 106: 745–779, 1995.
Dodt HU, Zieglgänsberger W. Infrared videomicroscopy: a new look at neuronal structure and function. Trends Neurosci 17: 453–458, 1994.[CrossRef][Web of Science][Medline]
Duch C, Levine RB. Changes in calcium signaling during postembryonic dendritic growth in Manduca sexta. J Neurophysiol 87: 1415–1425, 2002.
Ertel EA, Campbell KP, Harpold MM, Hofmann F, Mori Y, Perez-Reyes E, Schwartz A, Snutch TP, Tanabe T, Birnbaumer L, Tsien RW, Catterall WA. Nomenclature of voltage-gated calcium channels. Neuron 25: 533–535, 2000.[CrossRef][Web of Science][Medline]
Fedulova SA, Kostyuk PG, Veselovsky NS. Two types of calcium channels in the somatic membrane of new-born rat dorsal root ganglion neurones. J Physiol 359: 431–446, 1985.
Fox AP, Nowycky MC, Tsien RW. Kinetic and pharmacological properties distinguishing three types of calcium currents in chick sensory neurones. J Physiol 394: 149–172, 1987.
Gielow ML, Gu GG, Singh S. Resolution and pharmacological analysis of the voltage-dependent calcium channels of Drosophila larval muscles. J Neurosci 15: 6085–6093, 1995.[Abstract]
Gomora JC, Daud AN, Weiergraber M, Perez-Reyes E. Block of cloned human T-type calcium channels by succinimide antiepileptic drugs. Mol Pharmacol 60: 1121–1132, 2001.
Grolleau F, Lapied B. Two distinct low-voltage-activated Ca2+ currents contribute to the pacemaker mechanism in cockroach dorsal unpaired median neurons. J Neurophysiol 76: 963–976, 1996.
Grünewald B. Differential expression of voltage-sensitive K+ and Ca2+ currents in neurons of the honeybee olfactory pathway. J Exp Biol 206: 117–129, 2003.
Haag J, Borst A. Spatial distribution and characteristics of voltage-gated calcium signals within visual interneurons. J Neurophysiol 83: 1039–1051, 2000.
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. Pfluegers Arch 391: 85–100, 1981.[CrossRef][Web of Science][Medline]
Hayashi JH, Hildebrand JG. Insect olfactory neurons in vitro: morphological and physiological characterization of cells from the developing antennal lobes of Manduca sexta. J Neurosci 10: 848–859, 1990.[Abstract]
Hayashi JH, Levine RB. Calcium and potassium currents in leg motoneurons during postembryonic development in the hawkmoth Manduca sexta. J Exp Biol 171: 15–42, 1992.
Heidel E, Pflüger HJ. Ion currents and spiking properties of identified subtypes of locust octopaminergic dorsal unpaired median neurons. Eur J Neurosci 23: 1189–1206, 2006.[CrossRef][Web of Science][Medline]
Hess P, Lansman JB, Tsien RW. Calcium channel selectivity for divalent and monovalent cations. J Gen Physiol 88: 293–319, 1986.
Hildebrand JG, Shepherd GM. Mechanisms of olfactory discrimination: converging evidence for common principles across phyla. Annu Rev Neurosci 20: 595–631, 1997.[CrossRef][Web of Science][Medline]
Hille B. Ion Channels of Excitable Membranes. Sunderland, MA: Sinauer, 2001, p. 95–128.
Jeziorski MC, Greenberg RM, Anderson PA. The molecular biology of invertebrate voltage-gated Ca2+ channels. J Exp Biol 203: 841–856, 2000.[Abstract]
Johnson BD, Hockerman GH, Scheuer T, Catterall WA. Distinct effects of mutations in transmembrane segment IVS6 on block of L-type calcium channels by structurally similar phenylalkylamines. Mol Pharmacol 50: 1388–1400, 1996.[Abstract]
Johnson BR, Kloppenburg P, Harris-Warrick RM. Dopamine modulation of calcium currents in pyloric neurons of the lobster stomatogastric ganglion. J Neurophysiol 90: 631–643, 2003.
King GF. Modulation of insect Ca(v) channels by peptidic spider toxins. Toxicon 49: 513–530, 2007.[Medline]
Kirchhof BS, Mercer AR. Antennal lobe neurons of the honey bee, Apis mellifera, express a D2-like dopamine receptor in vitro. J Comp Neurol 383: 189–198, 1997.[CrossRef][Web of Science][Medline]
Kloppenburg P, Ferns D, Mercer AR. Serotonin enhances central olfactory neuron responses to female sex pheromone in the male sphinx moth Manduca sexta. J Neurosci 19: 8172–8181, 1999a.
Kloppenburg P, Hörner M. Voltage-activated currents in identified giant interneurons isolated from adult crickets Gryllus bimaculatus. J Exp Biol 201: 2529–2541, 1998.[Abstract]
Kloppenburg P, Kirchhof BS, Mercer AR. Voltage-activated currents from adult honeybee (Apis mellifera) antennal motor neurons recorded in vitro and in situ. J Neurophysiol 81: 39–48, 1999b.
Kloppenburg P, Zipfel WR, Webb WW, Harris-Warrick RM. Highly localized Ca2+ accumulation revealed by multiphoton microscopy in an identified motoneuron and its modulation by dopamine. J Neurosci 20: 2523–2533, 2000.
Kloppenburg P, Zipfel WR, Webb WW, Harris-Warrick RM. Heterogeneous effects of dopamine on highly localized voltage-induced Ca2+ accumulation in identified motoneurons. J Neurophysiol (August 29, 2007) doi:10.1152/jn.00660.2007.
Laurent G. A systems perspective on early olfactory coding. Science 286: 723–728, 1999.
Laurent G, Seymour-Laurent KJ, Johnson K. Dendritic excitability and a voltage-gated calcium current in locust nonspiking local interneurons. J Neurophysiol 69: 1484–1498, 1993.
Littleton JT, Ganetzky B. Ion channels and synaptic organization: analysis of the Drosophila genome. Neuron 26: 35–43, 2000.[CrossRef][Web of Science][Medline]
Mills JD, Pitman RM. Electrical properties of a cockroach motor neuron soma depend on different characteristics of individual Ca components. J Neurophysiol 78: 2455–2466, 1997.
Neher E. Correction for liquid junction potentials in patch clamp experiments. Methods Enzymol 207: 123–131, 1992.[Web of Science][Medline]
Patton C, Thompson S, Epel D. Some precautions in using chelators to buffer metals in biological solutions. Cell Calcium 35: 427–431, 2004.[CrossRef][Web of Science][Medline]
Pearson HA, Lees G, Wray D. Calcium-channel currents in neurons from locust (Schistocerca gregaria) thoracic ganglia. J Exp Biol 177: 201–221, 1993.[Abstract]
Saito M, Wu CF. Expression of ion channels and mutational effects in giant Drosophila neurons differentiated from cell division-arrested embryonic neuroblasts. J Neurosci 11: 2135–2150, 1991.[Abstract]
Schäfer S, Rosenboom H, Menzel R. Ionic currents of Kenyon cells from the mushroom body of the honeybee. J Neurosci 14: 4600–4612, 1994.[Abstract]
Strausfeld NJ, Hildebrand JG. Olfactory systems: common design, uncommon origins? Curr Opin Neurobiol 9: 634–639, 1999.[CrossRef][Web of Science][Medline]
Trequattrini C, Catacuzzeno L, Petris A, Franciolini F. Verapamil block of the delayed rectifier K current in chick embryo dorsal root ganglion neurons. Pfluegers Arch 435: 503–510, 1998.[CrossRef][Web of Science][Medline]
Triggle DJ. L-type calcium channels. Curr Pharm Des 12: 443–457, 2006.[CrossRef][Web of Science][Medline]
Wicher D, Penzlin H. Ca2+ currents in central insect neurons: electrophysiological and pharmacological properties. J Neurophysiol 77: 186–199, 1997.
Wicher D, Walther C, Wicher C. Non-synaptic ion channels in insects—basic properties of currents and their modulation in neurons and skeletal muscles. Prog Neurobiol 64: 431–525, 2001.[CrossRef][Web of Science][Medline]
Wilson RI, Mainen ZF. Early events in olfactory processing. Annu Rev Neurosci 29: 163–201, 2006.[CrossRef][Web of Science][Medline]
This article has been cited by other articles:
|
|
 |
|
  A. Husch, M. Paehler, D. Fusca, L. Paeger, and P. Kloppenburg Calcium Current Diversity in Physiologically Different Local Interneuron Types of the Antennal Lobe J. Neurosci., January 21, 2009; 29(3): 716 - 726. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
3 for all data points.
