© 1997 American Heart Association, Inc.
Articles |
Role of Ca2+ in Metabolic Inhibition–Induced Norepinephrine Release in Rat Brain Synaptosomes
Alfred and Baker Medical Unit, Baker Medical Research Institute and Alfred Hospital, Melbourne, Victoria, Australia.
Correspondence to Dr X.-J. Du, Alfred and Baker Medical Unit, Baker Medical Research Institute, Commercial Road, Prahran, Victoria 3181, Australia. E-mail xiao jun du@baker.edu.au
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Abstract |
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Ischemia and simulated ischemic conditions induce enhanced release of norepinephrine (NE) in the brain and the heart. Although studies with neuronal preparations demonstrated a rise in [Ca2+]i under energy-depleted conditions, such release of NE in the heart appears to be predominantly Ca2+ independent. Since Ca2+ overload occurs in ischemia or energy depletion and since a rise in [Ca2+]i triggers exocytosis without membrane depolarization, we tested the possibility, using brain synaptosomes, that increased NE release could be, at least in part, a consequence of raised [Ca2+]i. Brain synaptosomes were incubated with Krebs-Henseleit medium, and ischemia was mimicked by treatment with metabolic inhibitors. NE content in incubation medium (supernatant) and synaptosomes was analyzed chromatographically. Treatment with metabolic inhibitors reduced ATP content by 75% and increased [Ca2+]i by more than fourfold within minutes. Metabolic inhibition elicited NE release, which started within 10 minutes and reached a maximum after 30 minutes, with a corresponding 55% reduction in synaptosomal NE content after 40 minutes. NE release, together with a marked increase in [Ca2+]i, was also induced in energy-depleted synaptosomes by Ca2+ repletion after incubation with the Ca2+-free medium. Effects on NE release of various interventions to prevent Ca2+ overload were tested. Omission of Ca2+ from the incubation medium or loading synaptosomes with the Ca2+ chelator BAPTA-AM (20 and 100 µmol/L) prevented NE release, indicating a Ca2+-dependent mechanism. Inhibition of Ca2+ channels with
-conotoxin, cadmium, or nifedipine had no effect on NE release during energy depletion. In contrast, nickel and 3,4-dichlorobenzamil, Na+-Ca2+ exchange inhibitors, dose-dependently inhibited NE release. In conclusion, this study provides evidence that under energy-depleted conditions, Ca2+ overload in synaptosomes of noradrenergic neurons from the brain is an important mechanism for the enhanced release of NE and that a reversal of Na+-Ca2+ exchange may be the key pathway leading to intraneuronal Ca2+ overload.
Key Words: norepinephrine • metabolic inhibition • intracellular Ca2+ • synaptosome • Na+-Ca2+ exchange
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Introduction |
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Under normal physiological conditions, membrane depolarization of sympathetic neurons induces norepinephrine (NE) release by Ca2+-dependent exocytosis, with Ca2+ influx occurring predominantly through N-type Ca2+ channels.1 2
A large number of studies have documented the importance of Ca2+ overload in cellular injury caused by ischemia or energy depletion in neuronal and nonneuronal cells.3 4 5 6 7 Enhanced release of catecholamines occurs in the brain subjected to ischemia, hypoxia, and substrate deprivation.8 9 In vivo and in vitro studies using either synaptosomes, neurons, or chromaffin cells have demonstrated a rise in the intraneuronal Ca2+ ([Ca2+]i) during ischemia or energy depletion,5 10 11 12 13 and catecholamine release from chromaffin cells or PC12 cells has also been documented.6 10 12 In PC12 cells, the catecholamine release was abolished in a Ca2+-free medium.6 In chromaffin cells, although such release remains in Ca2+-free medium, the corelease of the intravesicular protein chromagranin A suggests an exocytotic mechanism.10 Furthermore, desipramine, which is potent in the suppression of NE release from the heart subjected to ischemia or anoxia, showed no effect on metabolic inhibition–mediated catecholamine release in chromaffin cells.10 However, Gustafson et al9 have demonstrated that NE release during brain ischemia in vivo, measured with a microdialysis technique, is partly inhibited by desipramine.
Under conditions of ischemia, anoxia, and metabolic inhibition, there is also an excessive and "spontaneous" release of NE from the heart.14 15 16 17 18 19 This release of NE has been attributed to a Ca2+-independent process, mediated by a reverse transport of NE by the neuronal uptake carrier (uptake1) as a result of raised [Na+]i and NE.15 16 17 20 In the perfused rat heart, a raised [Na+]i is presumably the consequence of a reduction in Na+ extrusion via Na+,K+-ATPase and enhanced Na+-H+ exchange activity.15 20 Interventions, which are expected to increase [Na+]i, enhance or accelerate such release.15 16 20 This "carrier-mediated efflux" can be inhibited by uptake1 inhibitors, such as desipramine and cocaine,14 15 16 17 and remains with a zero [Ca2+]0.14 15 16 20 The reasons for the apparent differences between neuronal preparations and the heart in the features of catecholamine release under ischemic conditions are still unclear.
It has been established that a rise in [Ca2+]i is capable of initiating exocytotic release of NE without membrane depolarization.21 22 This is in keeping with the finding from neuronal cells that Ca2+ influx via a reversal of Na+-Ca2+ exchange mediates massive release of catecholamines.23 24 Because an increase in [Ca2+]i usually follows a rise in [Na+]i, it is believed that reverse-mode Na+-Ca2+ exchange is involved.25 26
On the basis of these findings, we hypothesized that (1) Ca2+ influx and intraneuronal Ca2+ overload occur in neuronal cells under simulated ischemic conditions, (2) a reversal of Na+-Ca2+ exchange is an important cause of Ca2+ overload, and (3) under simulated ischemic conditions, a rise in [Ca2+]i in nerve varicosities, either from intraneuronal or extraneuronal sources, mediates the "spontaneous" release of NE by an exocytotic mechanism. This mechanism might contribute to enhanced catecholamine release in the heart and brain under energy-depleted conditions.
Synaptosome preparations are commonly used for studies involving neurotransmitter release and allow direct access to neuronal varicosities and to more effective interventions. There has been no detailed study with this preparation involving the ionic mechanisms of NE release under simulated ischemic conditions. Therefore, we have tested our hypotheses using incubated brain synaptosomes.
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Materials and Methods |
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Preparation of Synaptosomes
Sprague-Dawley rats (male; body weight, 250 to 300 g) were killed by decapitation. Synaptosomes were separated from whole brain, except cerebellum, using Ficoll-sucrose gradient centrifugation as described previously.27 28 All procedures were carried out at 4°C. Neuronal tissue was homogenized with a polytetrafluoroethylene-glass homogenizer in 0.32 mol/L sucrose containing ascorbic acid at 0.4 mmol/L and EDTA at 1.75 mmol/L (pH 7.4). This homogenate was spun at 1000g for 10 minutes, with the resulting supernatant centrifuged at 25 000g for 15 minutes. The pellet was washed once again with sucrose solution, resuspended in 5 mL of sucrose solution, and layered on a discontinuous two-step gradient containing Ficoll (dissolved in 0.32 mol/L sucrose) at 6.5% and 13%. Loaded gradients were placed in a swing-out rotor (SW-28, Beckman) and spun at 79 500g for 45 minutes. The interface band between 6.5% and 13% Ficoll was harvested, washed with 3 vol of the incubation medium (see below), and spun at 9900g for 10 minutes. The resultant pellet was resuspended in the medium and used for experiments immediately.
Incubation Procedure
The standard incubation medium was a Krebs-Henseleit solution containing (mmol/L) Na+ 148, K+ 4.0, Ca2+ 1.85, Mg2+ 1.05, HCO3- 25, PO43+ 0.5, EDTA 0.027, glucose 11, and ascorbic acid 0.4 and gassed with O2/CO2 (95%:5%, pH 7.4) before use. Incubation was carried out in microfuge tubes (with the lid on) in a water bath at 37°C with gentle shaking. Routinely, synaptosomes (
1 mg protein) were added to the incubation medium, with a total volume of 1.8 mL per tube. After a 30-minute preincubation period, vehicles or metabolic inhibitors (10 µmol/L rotenone and 1 mmol/L iodoacetate were used for most experiments) were added. Tubes were then harvested at 10, 20, 30, and 40 minutes, respectively, placed on ice, and centrifuged at 4°C. Supernatants were collected for catecholamine assay. Any chemical tested was added to the incubation tubes 10 minutes before the addition of metabolic inhibitors, except for BAPTA-AM.
Measurement of Neuronal Uptake of NE
As described previously,29 synaptosomes were incubated with L-[7-3H]NE (10 to 20 Ci/mmol; final concentration, 0.025 µCi/mL) for 30 minutes at 37°C in the presence of pargyline (100 µmol/L) and ascorbic acid (2 mmol/L). Two neuronal uptake inhibitors, desipramine and cocaine, were added to samples at various concentrations 10 minutes before the addition of [3H]NE. After a 30-minute incubation with [3H]NE, synaptosomes were washed three times with 2 mL of incubation medium at 37°C for 25 minutes each to remove loosely bound [3H]NE. The intrasynaptic NE was extracted by the addition of 1% Triton X-100 in a 5 mmol/L Tris base, and the radioactivity in the extract was counted with a scintillation counter. Nonspecific binding of [3H]NE was determined from the radioactivity in synaptosomes incubated at 4°C or in a medium with Na+ replaced by Li+ at 150 mmol/L.
Measurement of [Ca2+]i
Synaptosomes were incubated with Ca2+-free medium and loaded with fura 2-AM (5 µmol/L for 30 minutes at 30°C and then 5 minutes at 37°C). Loaded synaptosomes were washed with the Ca2+-free medium, centrifuged, and placed into a cuvette (1 mg protein in 3 mL) with continuous stirring, and fluorescence ratios were determined by excitation at 340/380 nm, alternatively, and recording emission at 505 nm with a Fluorolog-2 Spectrofluorometer (Spex Industries Inc), as previously described.30 Metabolic inhibitors or Ca2+ (final concentration, 1.85 mmol/L) was added sequentially into the cuvette either 3 minutes before or during the assay at the time specified below, and changes in fluorescence were monitored. As the addition of detergents or Ca2+ quenching agents changed the optical properties of the medium containing synaptosomes, autofluorescence was recorded at 340- and 380-nm excitations with identical aliquots of unloaded synaptosomes, and the values were subtracted from each excitation recording before calculation of Ca2+ values. Minimum and maximum fluorescence ratios were obtained by cell-free calibration using a solution mimicking the ionic composition of the cytosol. Autofluorescence values recorded at 380-nm excitation were <10% of the values observed in the presence of maximal [Ca2+]i (1.85 mmol/L; ionomycin, 2 µmol/L); this indicated that the level of unhydrolyzed fura 2 was minimal in synaptosomes. The extrasynaptosomal dye was estimated, in the presence of 1.85 mmol/L Ca2+, by separating the incubation medium and measuring fluorescence (340 nm), which is quenched after the addition of manganese.
NE and Other Biochemical Measurements
Supernatant from incubated synaptosomes was immediately frozen on dry ice and stored at -80°C until assay. NE in synaptosomes was extracted with 0.4 mol/L perchloric acid (containing 0.01% EDTA) and sonicated. NE and its metabolite dihydroxyphenylglycol (DHPG) were adsorbed with alumina and analyzed by HPLC with electrochemical detection.31 The interassay coefficient of variation was 5% for NE and 3% for DHPG.
Protein concentration was measured by Lowry's method with bovine albumin used as a standard. ATP in synaptosomes was extracted with a solution containing 7.5 mol/L KOH, 50 mmol/L KH2PO4, and 1 mol/L triethanolamine and quantified chromatographically, as previously described.32
Chemicals
Chemicals used and sources were as follows: Ficoll (Sigma Chemical Co), sucrose (ICN), iodoacetate (Sigma), rotenone (ICN), antimycin A (Sigma), 2-deoxy-D-glucose (Sigma), sodium cyanide (Sigma), desipramine (Sigma), cocaine (Sigma), lithium chloride (Sigma), nickel chloride (Ni2+, A.G. Darmstadt, E. Merck), cadmium chloride (Cd2+, BDH Ltd), veratridine (Sigma), -conotoxin (Peninsula Laboratories), pargyline (Sigma), A23187 (ICN), 3,4-dichlorobenzamil (provided by Dr E.J. Cragoe, Merck, Sharp and Dohme Laboratories, West Point, Pa), [3H]NE (New England Nuclear), fura 2-AM (Molecular Probes), BAPTA-AM (CalBiochem), and R56865 (Janssen Research Foundation). For chemicals dissolved in organic solvents (ethanol or dimethyl sulfoxide), final concentrations in the incubation medium were <0.1% for dimethyl sulfoxide and <0.05% for ethanol. Control preparations received the same concentrations of solvents. Neither of the agents was found to interfere with the catecholamine assay at the concentrations used.
Statistics
Results are presented as mean±SEM. Differences between and within groups were tested by one- or two-way ANOVA. The post hoc tests were unpaired or paired t tests. The statistically significant level was set at P<.05.
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Changes in Synaptosomal Content of ATP, NE, and DHPG by Metabolic Inhibitors
After a 30-minute preincubation (0 minute) with the standard medium, ATP content in synaptosomes was 3.19±0.07 nmol/mg protein and remained stable at 20 and 40 minutes of incubation with vehicles added (3.02±0.06 and 3.12±0.07 nmol/mg, respectively). Combined treatment with rotenone and iodoacetate (10 µmol/L and 1 mmol/L, respectively) reduced ATP to 0.76±0.05 and 0.84±0.03 nmol/mg (both P<.01 versus control).
Before incubation, the total content of NE in synaptosomes was 29.3±0.3 pmol/mg protein (ranging from 27.4 to 33.2 pmol/mg), and that of DHPG was 1.10±0.06 pmol/mg (n=22 from eight separate experiments). At the end of the 30-minute preincubation (ie, time 0 of the 40-minute incubation period) with the standard medium, NE content in synaptosomes was 20.6±0.5 pmol/mg protein and remained at this level throughout the next 40-minute incubation (20 minutes, 21.5±0.8 pmol/mg; 40 minutes, 21.0±0.7 pmol/mg; n=8 to 12 per group). This reduction in synaptosomal NE content was accompanied by an increase, in equal quantity, in the supernatant content of NE and DHPG (Fig 1A and 1B
). The DHPG content was 1.62±0.11 pmol/mg after the 30-minute period of preincubation and remained unchanged after 20 and 40 minutes (1.71±0.13 and 1.54±0.11 pmol/mg, respectively). As a result of the high background level of NE in the supernatant, NE release into the supernatant and effects of agents on such release were calculated from the difference in NE content of the supernatant per tube between vehicle-treated or metabolic inhibitor–treated samples harvested at the same time.
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Treatment with metabolic inhibitors depleted NE content in synaptosomes by 46% at 20 minutes (11.2±1.1 pmol/mg, n=8) and by 55% at 40 minutes (9.4±0.4 pmol/mg, n=8; both P<.01 versus respective controls). DHPG content was also significantly reduced after metabolic inhibition for 20 and 40 minutes (1.17±0.06 and 0.89±0.06 pmol/mg, respectively; P<.05 versus respective controls).
NE Release Induced by Metabolic Inhibition
In synaptosomes not treated with metabolic inhibitors, DHPG content in the supernatant increased by 3 pmol over the 40-minute incubation (Fig 1
). This was accompanied by a parallel decrease in NE content in the supernatant during this period (Fig 1A). Metabolic inhibition led to a further increase in the supernatant content of DHPG. However, this was accompanied by an increase in NE content in the supernatant (Fig 1A). Addition of the monoamine oxidase inhibitor pargyline (50 µmol/L) markedly reduced supernatant content of DHPG and substantially increased supernatant content of NE in synaptosomes without and with the addition of metabolic inhibitors. Fig 2A shows the time course of metabolic inhibition–induced NE release. In energy-depleted synaptosomes, NE release had already become significant within 10 minutes and increased progressively during the incubation period. Pargyline increased NE content in the supernatant by 1 to 2 pmol over the 10- to 40-minute period without a change in the time course of NE release. A similar release of NE was also induced by treatment with both 2-deoxyglucose (6 mmol/L) and antimycin A (1 µmol/L, Fig 2B). Sodium cyanide plus iodoacetate (1 mmol/L each) or rotenone alone (10 µmol/L) with no glucose for 40 minutes also induced NE release by 5.18±0.37 pmol and 6.78±0.87 pmol, respectively (four pairs of measurements per group).
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Ca2+ and Metabolic Inhibition–Induced NE Release
The role of Ca2+ in metabolic inhibition–mediated NE release was studied by omission of Ca2+ from the incubation medium or loading synaptosomes with the intracellular free Ca2+ chelator BAPTA.33 34 In synaptosomes incubated with Ca2+-free medium, NE content in the supernatant was slightly lower than that in the presence of Ca2+ at 1.85 mmol/L and decreased gradually over 40 minutes of incubation, with a rise in DHPG content (Fig 1C). NE release was induced by metabolic inhibition in the medium containing Ca2+ at 1.85 mmol/L but was abolished with a Ca2+-free medium, suggesting a mechanism dependent on the external Ca2+. Ca2+-free incubation largely inhibited NE release induced by a 40-minute incubation with cyanide plus iodoacetate or rotenone with no glucose (1.59±0.36 versus 5.18±0.27 pmol, and 1.46±0.19 versus 6.78±0.87 pmol; P<.05).
In another experiment carried out in the medium with 1.85 mmol/L Ca2+, synaptosomes were preincubated for 30 minutes with either vehicle or concentrations of BAPTA-AM at 2, 20, and 100 µmol/L, respectively, and then treated with metabolic inhibitors for 40 minutes. NE release was found to be dose-dependently inhibited by BAPTA, with an 80% inhibition achieved at 100 µmol/L (Fig 3).
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Because Ca2+ influx could occur through Ca2+ channels or a reversal of Na+-Ca2+ exchange, we tested the importance of these Ca2+ entry pathways during metabolic inhibition. In this series of experiments, -conotoxin (50 nmol/L), Cd2+ (0.1 mmol/L), and nifedipine (0.1 µmol/L) were added 10 minutes before the addition of metabolic inhibitors. NE release by metabolic inhibition was observed in all vehicle-treated samples, and treatment with the three agents showed no effect on such NE release (Table), excluding a role for Ca2+ influx through neuronal Ca2+ channels. In further experiments, the effects of Ni2+ and 3,4-dichlorobenzamil, inhibitors of the Na+-Ca2+ exchanger,35 36 37 38 39 on NE release during metabolic inhibition were examined. Treatment with either Ni2+ (0.1 and 1.0 mmol/L) or 3,4-dichlorobenzamil (3 and 30 µmol/L) partly and largely suppressed NE release (Fig 4A and 4B). Under our experimental conditions, R56865 at 1 and 10 µmol/L dose-dependently inhibited NE release mediated by metabolic inhibition (Fig 4C).
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We also tested the role of Ca2+ in NE release in synaptosomes treated with metabolic inhibitors using a different protocol. Synaptosomes were incubated in Ca2+-free medium for 35 minutes in the presence of metabolic inhibitors. Ca2+ was then added to the medium, and samples were harvested 5 minutes after the addition. Readmission of Ca2+ induced an increase in NE content in the supernatant over respective control levels (20 minutes, 1.76±0.28 pmol; 40 minutes, 2.26±0.14 pmol). In the samples without treatment with metabolic inhibitors, Ca2+ repletion, after incubation with Ca2+-free medium for 40 minutes, did not induce NE release (0.17±0.38 pmol, P<.01 versus metabolic inhibition).
Effect of Desipramine and Cocaine on Metabolic Inhibition–Induced NE Release
In energy-depleted synaptosomes, treatment with desipramine at 0.3 µmol/L inhibited NE release by 75% to 90% (Fig 5). For comparison, the effect of another neuronal uptake inhibitor, cocaine, on metabolic inhibition–induced NE release was tested. Synaptosomes were pretreated with vehicle or cocaine at 10, 30, and 100 mmol/L, respectively, 10 minutes before the addition of rotenone and iodoacetate. Pargyline was present at 50 µmol/L. After metabolic inhibition for 40 minutes, the increase in NE content in the supernatant was 9.2±0.3 pmol (n=4) in control samples. Cocaine at 10 and 30 µmol/L showed no significant effect on NE release by metabolic inhibition (8.2±0.3 and 7.9±0.6 pmol, respectively; four measurements each; Fig 5B). At 100 µmol/L, a concentration that inhibited [3H]NE uptake by 90%, cocaine reduced NE release by 23% (6.8±0.2 pmol, P<.05 versus control, Fig 5B). The ineffectiveness of cocaine was also observed in another experiment in the absence of pargyline (5.7±0.7 pmol versus 6.9±0.5 pmol; n=6 and 8, respectively).
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In synaptosomes with metabolic inhibition, desipramine totally prevented Ca2+ readmission–induced increase in supernatant NE content compared with respective control levels (20 minutes, -0.16±0.36 versus 1.76±0.28 pmol; 40 minutes, -0.08±0.10 versus 2.26±0.14 pmol; both P<.01).
Changes in [Ca2+]i
Changes in [Ca2+]i by metabolic inhibition were examined in fura 2–loaded synaptosomes. Ca2+ (final concentration, 1.85 mmol/L) and metabolic inhibitors were then added into the cuvette after equilibration. [Ca2+]i was 260±8 nmol/L (n=17) in the Ca2+-free medium and increased to 499±52 nmol/L (n=8) in the medium with Ca2+ at 1.85 mmol/L. Addition of metabolic inhibitors induced a marked rise in [Ca2+]i by 1 µmol/L after 4 minutes (Fig 6A). In Ca2+-free medium, metabolic inhibition induced a slow increase in [Ca2+]i from 252±6 to 377±8 nmol/L in 5 minutes (P<.01). In synaptosomes treated with metabolic inhibitors and incubated in Ca2+-free medium, reintroduction of Ca2+ led to a large rise in [Ca2+]i (Fig 6B). This rise in [Ca2+]i appeared to be biphasic, with an initial rapid phase of [Ca2+]i increase by 600 nmol/L in 30 seconds, followed by a slowly rising [Ca2+]i at 60 nmol/L per minute over the experimental period.
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We examined the possibility that metabolic inhibition causes leakage of fura 2, which contributes to the rise in [Ca2+]i. The fluorescence intensity from extrasynaptosomal fura 2 at 340 nm was 37078±856 cps in control samples (n=3), contributing to 10.6±0.6% of total fluorescence in the presence of 1.85 mmol/L Ca2+. There was no significant increase in the fluorescence intensity 7 minutes after metabolic inhibition (40 939±10 231 cps, n=6).
Using these protocols, we found that desipramine at 0.3 µmol/L had no effect on the rise in [Ca2+]i by ATP depletion (data not shown). Nickel (1 mmol/L) largely suppressed the rise in [Ca2+]i induced by Ca2+ readmission and by metabolic inhibition (Fig 7). The effect of dichlorobenzamil on [Ca2+]i could not be studied because of a strong interference on fura 2 fluorescence by this agent.
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NE Uptake and Effect of Desipramine and Cocaine
When incubated with the standard medium, synaptosomes exhibited the capability to accumulate [3H]NE, with synaptosomal radioactivity of 7763±297 dpm/mg protein (16 measurements from four separate experiments). This uptake of NE was inhibited, in a dose-dependent manner, by both desipramine (percentage of control, 61±1% at 0.1 µmol/L, 51±2% at 0.3 µmol/L, 35±4% at 3 µmol/L, and 29±2% at 30 µmol/L; 6 to 11 measurements per dosage) and cocaine (52±1% at 10 µmol/L, 26±1% at 30 µmol/L, and 9±1% at 100 µmol/L; 4 to 13 measurements per dosage, Fig 4B).
Evoked NE Release
In synaptosomes incubated with the standard medium, several agents, veratridine (voltage-gated Na+ channel opener, 10 µmol/L, n=12), A23187 (Ca2+ ionophore, 5 µmol/L, n=3), and K+ (80 mmol/L, n=5), increased NE content in the supernatant by 3.62±0.25, 1.85±0.09, and 3.21±0.21 pmol, respectively, suggesting induced release. NE release evoked by veratridine was inhibited by 80% to 95% with Cd2+ (0.1 mmol/L), Ni2+ (1 mmol/L), BAPTA-AM (20 µmol/L), and Ca2+ omission (results not shown), indicating a Ca2+-dependent exocytotic mechanism.
Veratridine (10 µmol/L)–evoked NE release was examined in synaptosomes incubated with the standard medium (n=4) and in synaptosomes treated with metabolic inhibitors for 10, 20, 30, and 40 minutes (four measurements each). In energy-depleted synaptosomes, NE release in response to veratridine was suppressed at all time points tested (from 5.18±0.29 to 0.41±0.23, -0.49±0.44, 0.60±0.32, and 0.80±0.12 pmol; P<.01).
In this preparation, incubation for 40 minutes in a medium with Na+ replaced by an equimolar concentration of LiCl led to NE release (5.63±0.29, n=4). NE release by Na+-free incubation was largely inhibited by Ni2+ at 1 mmol/L to 1.82±0.26 pmol/L (P<.05).
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Discussion |
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The major goal of the present study was to examine whether NE release from brain synaptosomes under conditions of metabolic inhibition was mediated by a Ca2+-dependent or Ca2+-independent mechanism. Several major observations have been made: (1) metabolic inhibition substantially reduced the synaptosomal content of ATP and NE and enhanced NE release; (2) metabolic inhibition led to a marked rise in [Ca2+]i in the presence of external Ca2+; (3) interventions aimed at preventing the rise in [Ca2+]i by metabolic inhibition, such as BAPTA loading and Ca2+ removal, essentially abolished NE release; (4) Ni2+ and dichlorobenzamil inhibited this NE release; and (5) desipramine suppressed NE release induced by ATP depletion without affecting the rise in [Ca2+]i, whereas cocaine was ineffective.
Under our experimental conditions, incubated synaptosomes maintained function, as evidenced by the capability of uptake and storage of NE, release of NE in response to various stimuli, and maintenance of [Ca2+]i at a low level. Energy depletion induced a marked increase in [Ca2+]i and release of NE into the supernatant, together with a partial depletion of synaptosomal NE content. In this model, synaptosomes were contaminated by free mitochondrial particles28 ; thus, monoamine oxidase activity was present outside as well as inside the synaptosome. Because most experiments were performed in the absence of pargyline, the actual quantity of NE released by metabolic inhibition would have been higher than that measured in the presence of pargyline because of the metabolism of NE by this enzyme. Furthermore, the presence of monoamine oxidase activity outside synaptosomes confounded the interpretation of DHPG data.
Under physiological conditions, NE release in response to nerve activation is via a Ca2+-dependent exocytotic mechanism. In the perfused rat heart subjected to ischemia, anoxia (without substrate), or metabolic inhibition, a locally mediated and nerve activity–independent NE release occurs.14 15 Two features distinguish this release of NE from that occurring in response to neural activation: it occurs in the absence of external Ca2+, and it is suppressed by uptake1 inhibitors, such as desipramine and cocaine.14 15 16 17 Thus, such local NE release has been considered to be a process mediated by the neuronal uptake carrier transporting NE in the outward direction.14 15 16 17 This occurs as a two-stage process. NE leaks from storage vesicles into the neuroplasma because of a collapse of the transvesicular pH gradient, which is energy dependent and is normally responsible for retention of NE within vesicles.20 Free NE in the axoplasma is protected from metabolism by monoamine oxidase because of an insufficient oxygen supply.20 There is also a rise in [Na+]i as a result of the suppression of K+,Na+-ATPase and increase in Na+-H+ exchange. As a result of elevated [Na+]i and axoplasmic NE, the uptake carrier mediates the outward transport of NE, leading to NE efflux.14 15 16 17 20 Pharmacological manipulations that are expected to increase [Na+]i and to induce NE leakage from vesicles can mimic ischemic or anoxic NE efflux.15 16 20 This explanation fits the fact that uptake1 inhibitors and agents, which prevent intracellular Na+ overload, suppress such release.14 17 20
In the present study with brain synaptosomes, however, several observations do not reconcile with carrier-mediated efflux as the primary mechanism responsible for NE release. Ca2+ omission and chelation of intracellular Ca2+ with BAPTA largely inhibited NE release induced by metabolic inhibition, providing evidence that under our experimental conditions, such NE release is dependent on Ca2+ influx and a substantial rise in [Ca2+]i. Our finding is in keeping with a report that cyanide-induced catecholamine release in PC12 cells was abolished in Ca2+-free medium.6 Further support for this notion is the finding that Ca2+ readmission to energy-depleted synaptosomes, incubated in the Ca2+-free medium, induced NE release. NE release by this protocol was accompanied by a marked rise in [Ca2+]i in synaptosomes after metabolic inhibition, a finding similar to previous studies with various neuronal preparations.10 11 40 In addition, the uptake1 inhibitor, cocaine, is largely ineffective in the inhibition of energy depletion–induced NE release at doses (10 to 100 µmol/L) that inhibited NE uptake by 50% to 90%.
According to the hypothesis of carrier-mediated efflux, NE first accumulates in the axoplasma and, when oxygen is available, is subjected to oxidation by monoamine oxidase to form DHPG, a process independent of ATP supply. In the perfused rat heart with an intact activity of monoamine oxidase, overflow of DHPG with little NE efflux occurs when leakage of NE from vesicles and a reduction in Na+ gradient across neuronal membrane are induced using pharmacological approaches.15 20 Furthermore, in perfused rat hearts with metabolic inhibition but in the presence of oxygen, overflow of DHPG proceeds that of NE by 10 minutes.15 In the present study with synaptosomes in which ATP synthesis was prevented but oxygen was still present, NE release induced by metabolic inhibition was evident, and treatment with pargyline led to a 1- to 2-pmol increase in NE content in the supernatant. This finding does not suggest a significant leakage of NE from storage vesicles into the axoplasma during metabolic inhibition. Indeed, we found a reduction, not an increase, in synaptosomal content of DHPG after metabolic inhibition. However, energy depletion–induced NE release was inhibited by 26% with 100 µmol/L cocaine, a dose that almost totally suppressed [3H]NE uptake (Fig 5). Thus, the carrier-mediated and Ca2+-independent mechanism, which plays a major role in ischemic and energy depletion–induced NE release in the heart,5 6 7 8 9 10 11 12 13 14 15 16 17 20 may contribute to overall NE release from energy-depleted brain synaptosomes.
Several mechanisms might account for elevated [Ca2+]i during metabolic inhibition, such as Ca2+ influx through Ca2+ channels, Ca2+ entry through the reversal of Na+-Ca2+ exchange, and mobilization of intracellular Ca2+. In the present study, a significant rise in [Ca2+]i is also observed, after metabolic inhibition, in synaptosomes incubated with Ca2+-free medium. However, this rise is much smaller than that in the presence of external Ca2+ and is not associated with significant NE release. Therefore, it seems that in energy-depleted synaptosomes, a certain threshold level of [Ca2+]i is necessary to induce exocytosis and that Ca2+ influx is required to reach this level. Both Ni2+ and dichlorobenzamil possess inhibitory effects on the Na+-Ca2+ exchanger that exists in neuronal cells23 24 41 and are effective in the suppression of such NE release. Ni2+ was also potent in the suppression of NE release induced by Na+-free incubation (to induce Ca2+ influx via reverse-mode Na+-Ca2+ exchange) and the rise in [Ca2+]i in energy-depleted synaptosomes. Na+-Ca2+ exchange is voltage dependent. Depolarization may occur during ATP depletion because of the inhibition of Na+,K+-ATPase and K+ efflux, and this would favor Ca2+ influx through increased Na+-Ca2+ exchange. Studies have shown that in neuronal and nonneuronal tissues exposed to ischemic, anoxic, or energy-depleted conditions, a rise in [Ca2+]i is the consequence of reverse-mode Na+-Ca2+ exchange or mobilization of intracellular Ca2+ stores.4 23 24 42 43 In the present study, a role for reverse-mode Na+-Ca2+ exchange in Ca2+ overload is suggested by the inhibition of metabolic inhibition–induced NE release with both Ca2+ removal and Na+-Ca2+ exchange inhibitors. Inasmuch as elevated [Ca2+]i in nerve varicosities may itself induce vesicular exocytosis,21 22 23 24 this Ca2+ overload following energy depletion may mediate NE release independent of membrane depolarization and Ca2+ influx through voltage-gated Ca2+ channels. Indeed, Ca2+-free or pharmacological inhibition of the Na+-Ca2+ exchanger significantly protected nerves from anoxic damage.4 43 44 In addition, raised [Ca2+]i may stimulate Na+-H+ exchange, leading to a further elevation of [Na+]i.45 The latter can then enhance the influx of Ca2+ through Na+-Ca2+ exchange.37 Contributing factors to the increased [Ca2+]i include Ca2+-induced Ca2+ release and a reduced Ca2+ extrusion and sequestration via endoplasmic reticulum Ca2+-ATPase.46
It is well known that ATP depletion can inhibit Na+-Ca2+ exchange.47 48 However, the inhibition of NE release by Ni2+ and dichlorobenzamil suggests a maintained operation of the exchanger in the reverse mode. In addition to ATP supply, the operation of the exchanger and the direction of Ca2+ movement are controlled by several additional factors, including membrane potential and transmembrane Na+ and Ca2+ gradients. Under energy-depleted conditions, there is an occurrence of membrane depolarization and Na+ overload that might be sufficient to overcome the inhibition by reduced ATP levels on the reverse-mode exchange. This has been suggested by several studies.25 47
The inhibitory action of Ni2+ on Na+-Ca2+ exchange has been well defined.36 37 38 However, this ion can also inhibit a number of voltage- or receptor-operated ionic channels for Na+, Ca2+, K+, and Cl-.35 Although our data do not suggest a role for Ca2+ channels in mediating Ca2+ overload, some of these additional actions of Ni2+ might contribute to the inhibition of NE release in energy-depleted synaptosomes. For example, blockade of Na+ channels may alleviate Na+ overload, which, in turn, enhances Ca2+ influx through reverse-mode Na+-Ca2+ exchange.4 25 One study has shown that Ni2+ entry into cells, via divalent ionic channels, is very limited.49 Thus, a contribution by intrasynaptic actions of Ni2+ is unlikely.
Protection by R56865 against Na+ and Ca2+ overload during ischemia, reperfusion, and hypoxia has been well documented.25 50 51 It is believed that the main mechanism for this protection involves blockade of voltage-gated Na+ channels.25 50 52 The inhibition by R56865 of metabolic inhibition–induced NE release suggests that Na+ overload during energy depletion is partly due to Na+ influx via Na+ channels.
In the present study, desipramine was effective in the inhibition of NE release induced by metabolic inhibition, which, as discussed, is by a Ca2+-dependent mechanism. It seems difficult to attribute this effect of desipramine to its inhibition of neuronal uptake, because another uptake1 inhibitor, cocaine, was largely ineffective. Desipramine also suppressed Ca2+ repletion–induced NE release after a period of metabolic inhibition and Ca2+-free incubation. When this protocol was used, readmission of Ca2+ induced Ca2+ influx, most probably in exchange for Na+ already accumulated in the synaptosomes. Studies with neural preparations have observed that tricyclic antidepressants, including desipramine, suppress exocytotic NE release53 54 and block Ca2+ entry via various pathways, including voltage-gated Ca2+ channels, N-methyl-D-aspartate receptor–gated Ca2+ channels, and reverse-mode Na+-Ca2+ exchange.55 56 57 However, these effects require high concentrations under physiological conditions (10 to 50 µmol/L). In the present study, we did not observe any suppression by desipramine on the rise in [Ca2+]i during metabolic inhibition. Recent studies with neuronal preparations have shown that desipramine may inhibit neuronal signal transduction pathways involving inositol triphosphate, protein kinase C, and adenylyl cyclase–cAMP.58 59 60 Thus, the mechanism of desipramine remains to be investigated and might be involved in other processes mediating NE release under energy-depleted conditions.
NE release induced by a Na+ channel opener, veratridine, was found to be largely suppressed during 10- to 40-minute periods of metabolic inhibition. This result is in keeping with two previous studies with chromaffin cells showing a reduced exocytotic release of NE by energy depletion.10 12 Part of the reason for this suppressed releasing response in energy-depleted synaptosomes is reduced synaptosomal content of NE. In contrast, in energy-depleted hearts from rat and guinea pig, but not in human atrial tissue, NE release evoked by direct membrane depolarization or nicotine stimulation is substantially enhanced during the early stages (10 to 30 minutes).18 61 62 63 64 Such exaggerated exocytosis occurs when the carrier-mediated NE efflux is evident, and it has been postulated that Ca2+ overload with elevated [Ca2+]i potentiated NE exocytotic release.18 61 Although nerve stimulation–evoked NE release was attenuated progressively within the first 20 minutes,18 62 63 64 this may be partly due to an impairment of neuronal conduction under ischemic conditions.65
The between-model differences in features of NE release are interesting and require further discussion. In hearts exposed to ischemia or substrate-free anoxia, NE release is maintained in the absence of external Ca2+, demonstrating evidence of a mechanism independent of Ca2+ influx.14 16 20 In the present study, metabolic inhibition–induced NE release from synaptosomes is dependent on external Ca2+. Several factors might be responsible. Differences between central and peripheral adrenergic synaptosomes in functional properties cannot be excluded. Wakade and colleagues29 66 have demonstrated substantial changes in NE-releasing properties of adrenergic neurons after coculture with cardiac myocytes. They also found, in cultured sympathetic neurons, that Ca2+ influx through reverse-mode Na+-Ca2+ exchange triggered massive release of NE and that this release was attenuated 80% when neurons were cocultured with cardiac myocytes.24 Coculture of neurons with cardiac myocytes also enhanced neuronal uptake activity.29 Thus, further studies are required to test whether neuromodulatory effects of cardiac cells are responsible for the differences between heart models and brain synaptosomes in the mechanisms of NE release under ischemic and simulated ischemic conditions.
Several limitations in the present study should be addressed. First, although we examined NE release mechanisms in relation to changes in [Ca2+]i, this preparation contains synaptosomes from a variety of neurons using different transmitters. We cannot exclude the possibility that such heterogeneous synaptosomes might interfere with the end points studied. However, findings from the experiment with BAPTA indicate the effectiveness of loading adrenergic synaptosomes with fura 2, a derivative from BAPTA with a similar structure. Furthermore, several interventions (Ca2+ free, Ca2+ readmission, and Ni2+) changed NE release and [Ca2+]i in the same direction, indicating that these measurements reflect changes in adrenergic synaptosomes. Second, the intraneuronal mechanisms by which desipramine suppresses NE release by energy depletion remains unanswered. Third, the main goal of the present study was to define whether the mechanism responsible for NE release is Ca2+ dependent. Although the results support the view that a reverse-mode Na+-Ca2+ exchange is important in mediating Ca2+ overload, [Na+]i was not measured, and the pathways leading to Na+ overload are less clearly defined.
In conclusion, the present study provides evidence that in brain synaptosomes metabolic inhibition–induced NE release is mainly via a Ca2+-dependent mechanism and that a reversal of Na+-Ca2+ exchange may play an important role in mediating Ca2+ overload. The same mechanism may account, largely or in part, for the enhanced NE release from the brain and the heart under conditions of ischemia and energy depletion, although a mechanism independent of Ca2+ influx plays a major role in NE release from the heart under these conditions.
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Acknowledgments |
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This study was supported by a grant from National Heart Foundation of Australia (G93M3888) and an institute grant to the Baker Medical Research Institute from the National Health and Medical Research Council of Australia. We thank Professor James A. Angus (Department of Pharmacology, University of Melbourne) for his helpful suggestions and supply of
-conotoxin. A sample of 3,4-dichlorobenzamil was kindly supplied by Dr Edward J. Cragoe, Jr, and prepared by the method described by Cragoe et al.67 A test sample of R56865 was generously supplied by Dr Marcel Janssen (Janssen Research Foundation, Belgium). We thank Helen Cox and Andrea Turner for their help in the catecholamine assay and Dr Steve Richards for his help in the ATP assay. The kind help of Dr E.A. Woodcock, who provided laboratory facilities, is gratefully acknowledged. Received June 6, 1996; accepted October 28, 1996.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
1. Hirning LD, Fox AP, McCleskey EW, Olivera BM, Thayer SA, Miller RJ, Tsien RW. Dominant role of N-type Ca2+ channels in evoked release of norepinephrine from synaptic neurons. Science. 1988;239:58-61.
2. Daniell LC, Leslie SW. Correlation of rates of calcium entry and release of endogenous norepinephrine in rat brain region synaptosomes. J Neurochem.. 1986;46:249-256.[Medline] [Order article via Infotrieve]
3. Choi DW. Calcium-mediated neurotoxicity: relationship to specific channel types and role in ischemic damage. Trends Neurosci.. 1988;11:465-469.[Medline] [Order article via Infotrieve]
4.
Stys PK, Ransom BR, Waxman SG, Davis PK. Role of extracellular calcium in anoxic injury of mammalian central white matter. Proc Natl Acad Sci U S A.. 1990;87:4212-4216.
5. Uematsu D, Greenberg JH, Reivich M, Karp A. In vivo measurement of cytosolic free calcium during cerebral ischemia and reperfusion. Ann Neurol.. 1988;24:420-428.[Medline] [Order article via Infotrieve]
6. Kanthasamy AG, Maduh EU, Peoples RW, Borowitz IJ, Isome GE. Calcium mediation of cyanide-induced catecholamine release: implications for neurotoxicity. Toxicol Appl Pharmacol.. 1991;110:271-282.
7.
Steenbergen C, Murphy E, Watts JA, London RE. Correlation between cytosolic free calcium, contracture, ATP, and irreversible ischemic injury in perfused rat heart. Circ Res.. 1990;66:135-146.
8. Damsma G, Boisvert DP, Mudrick LA, Wenkstern D, Fibiger HC. Effects of transient forebrain ischemia and pargyline on extracellular concentrations of dopamine, serotonin, and their metabolites in the rat striatum as determined by in vivo microdialysis. J Neurochem.. 1990;54:801-808.[Medline] [Order article via Infotrieve]
9. Gustafson I, Westerberg EJ, Wieloch T. Extracellular brain cortical levels of noradrenaline in ischemia: effects of desipramine and post-ischemic administration of idazoxan. Exp Brain Res.. 1991;86:555-561.[Medline] [Order article via Infotrieve]
10.
Dry KL, Phillips JH, Dart AM. Catecholamine release from bovine adrenal chromaffin cells during anoxia or metabolic inhibition. Circ Res.. 1991;69:466-474.
11.
Nachshen DA. Regulation of cytosolic calcium concentration in presynaptic nerve endings isolated from rat brain. J Physiol (Lond).. 1985;363:87-101.
12. Itoh A, Miwa S, Koshimura K, Akiyama Y, Takagi Y, Yamagata S, Kikuchi H, Masaki T. Ischemia-induced changes in catecholamine release and their mechanisms: a study using cultured bovine adrenal chromaffin cells. Brain Res.. 1994;643:266-275.[Medline] [Order article via Infotrieve]
13. Carroll JM, Toral-Barza L, Gibson G. Cytosolic free calcium and gene expression during chemical hypoxia. J Neurochem.. 1992;59:1836-1843.[Medline] [Order article via Infotrieve]
14.
Schomig A, Dart AM, Dietz R, Mayer E, Kubler W. Release of endogenous catecholamines in the ischemic myocardium of the rat, A: locally mediated release. Circ Res.. 1984;55:689-701.
15.
Schomig A, Kurz T, Richardt G, Schomig E. Nonexocytotic release of endogenous noradrenaline in the ischemic and anoxic heart: mechanism and metabolic requirements. Circ Res.. 1987;60:194-205.
16. Dart AM, Riemersma RA, Schomig A, Ungar A. Metabolic requirements for release of endogenous noradrenaline during myocardial ischaemia and anoxia. Br J Pharmacol.. 1987;90:43-50.[Medline] [Order article via Infotrieve]
17.
Carlsson L, Abrahamsson T. Characterization of the inhibitory effect of some antidepressant drugs on the outward transport of norepinephrine in the ischemic myocardium J Pharmacol Exp Ther. 1988;247:715-720.
18.
Du X-J, Dart AM. Mechanisms of noradrenaline release in the anoxic heart of the rat. Cardiovasc Res. 1993;27:2011-2015.
19.
Akiyama T, Yamazaki T, Ninomiya I. Differential regional responses of myocardial interstitial noradrenaline levels to coronary occlusion. Cardiovasc Res.. 1993;27:817-822.
20.
Schomig A, Kurz T, Richardt G, Schomig E. Neuronal sodium homeostasis and axoplasmic amine concentration determine calcium-independent noradrenaline release in normoxic and ischemic rat heart. Circ Res.. 1988;63:214-226.
21. Zucker RS, Hayden PG. Membrane potential has no direct role in evoking neurotransmitter release. Nature. 1988;335:360-362.[Medline] [Order article via Infotrieve]
22. Mulkey RM, Zucker RS. Action potentials must admit calcium to evoke transmitter release. Nature. 1991;350:153-155.[Medline] [Order article via Infotrieve]
23. Wakade AR, Przywara DA, Bhave SV, Chowdhury PS, Bhave A, Wakade TD. Massive exocytosis triggered by sodium-calcium exchange in sympathetic neurons is attenuated by co-culture with cardiac cells. Neuroscience. 1993;55:813-821.[Medline] [Order article via Infotrieve]
24. Torok TL. Norepinephrine and catecholamine release from peripheral sympathetic nerves and chromaffin cells maintained in primary tissue culture: the role of sodium-calcium exchange. Ann N Y Acad Sci.. 1991;693:631-641.
25.
Haigney MCP, Miyata L, Lakatta EG, Stern MD, Silverman HS. Dependence of hypoxic cellular calcium loading on Na+-Ca2+ exchange. Circ Res.. 1992;71:547-557.
26.
Jelicks LA, Gupta RK. Multinuclear NMR studies of the Langendorff perfused rat heart. J Biol Chem.. 1989;264:15230-15235.
27. Wolf WA, Bobik A. Effects of 5,6-dihydroxytryptamine on the release, synthesis, and storage of serotonin: studies using rat brain synaptosomes. J Neurochem.. 1988;50:534-542.[Medline] [Order article via Infotrieve]
28. Cotman CW, Matthews DA. Synaptic plasma membranes from rat brain synaptosomes: isolation and partial characterization. Biochem Biophys Acta.. 1971;249:380-394.[Medline] [Order article via Infotrieve]
29. Wakade AR, Bhave SV. Facilitation of noradrenergic character of sympathetic neurons by co-culturing with heart cells. Brain Res.. 1988;458:115-122.[Medline] [Order article via Infotrieve]
30.
Neylon CB, Nickashin A, Little PJ, Tkachuk VA, Bobik A. Thrombin-induced Ca2+ mobilization in vascular smooth muscle utilizes a slowly ribosylating pertussis toxin-sensitive G protein. J Biol Chem.. 1992;267:7295-7302.
31. Medvedev OS, Esler MD, Angus JA, Cox HS, Eisenhofer G. Simultaneous determination of plasma noradrenaline and adrenaline kinetics. Naunyn Schmiedebergs Arch Pharmacol.. 1990;341:192-199.[Medline] [Order article via Infotrieve]
32. Olivares J, Verdys M. Isocratic high-performance liquid chromatographic method for studying the metabolism of blood plasma pyrimidine nucleosides and bases: concentration and radioactivity measurements. J Chromatogr.. 1988;434:111-121.[Medline] [Order article via Infotrieve]
33. Tsien RW. New calcium indicators and the buffers with high selectivity against magnesium and protons: design, synthesis, and properties of prototype structure. Biochemistry.. 1980;19:2396-2404.[Medline] [Order article via Infotrieve]
34. Fredholm BB, Hu PS. Effect of an intracellular calcium chelator on the regulation of electrically evoked [3H]-noradrenaline release from rat hippocampal slices. Br J Pharmacol.. 1993;108:126-131.[Medline] [Order article via Infotrieve]
35. Kiss T, Osipenko ON. Toxic effects of heavy metals on ionic channels. Pharmacol Rev.. 1994;46:245-267.[Medline] [Order article via Infotrieve]
36.
Schulze P, Kofuji P, Hadley R, Kirby MS, Kieval RS, Doering A, Niggli E, Lederer WJ. Sodium/calcium exchanger in heart muscle: molecular biology, cellular function, and its special role in excitation-contraction coupling. Cardiovasc Res.. 1993;27:1726-1734.
37.
Sham JS, Cleemann L, Morad M. Gating of the cardiac Ca2+ release channel: the role of Na+ current and Na+-Ca2+ exchange. Science. 1992;255:850-853.
38.
Kimura J, Miyamae S, Noma A. Identification of sodium-calcium exchange current in single ventricular cells of guinea-pig. J Physiol (Lond).. 1987;384:199-222.
39. Slaughter RS, Garcia ML, Cragoe EJ Jr, Reeves JP, Kaczorowski GJ. Inhibition of sodium-calcium exchange in cardiac sarcolemmal membrane vesicles, 1: mechanism of inhibition by amiloride analogues. Biochemistry. 1988;27:2403-2409.[Medline] [Order article via Infotrieve]
40. Kauppinen RA, McMahon HT, Nicholls DG. Ca2+-dependent and Ca2+-independent glutamate release, energy status and cytosolic free Ca2+ concentration in isolated nerve terminals following metabolic inhibition: possible relevance to hypoglycaemia and anoxia. Neuroscience. 1988;27:175-182.[Medline] [Order article via Infotrieve]
41. Blaustein MP, Goldman WF, Fontane G, Krueger BK, Santiago EM, Steele TD, Weiss DN, Yarowsky PJ. Physiological roles of the sodium-calcium exchanger in nerve and muscle. Ann N Y Acad Sci.. 1991;639:255-273.
42. Morris AC, Hagler HK, Willerson JT, Buja LM. Relationship between calcium loading and impaired energy metabolism during Na+, K+ pump inhibition and metabolic inhibition in cultured neonatal rat cardiac myocytes. J Clin Invest.. 1989;83:1876-1887.
43. Stys PK, Waxman SG, Ransom BR. Na+-Ca2+ exchanger mediates Ca2+ influx during anoxia in mammalian central nervous system white matter. Ann Neurol.. 1991;30:375-380.[Medline] [Order article via Infotrieve]
44. Siesjo BK. Calcium in the brain under physiological and pathological conditions. Eur Neurol. 1990;30(suppl 2):3-9.
45.
Little PJ, Weissberg PL, Cragoe EJ, Bobik A. Dependence of Na+/H+ antiport activation in cultured rat aortic smooth muscle on calmodulin, calcium, and ATP. J Biol Chem.. 1988;263:16780-16786.
46. Simpson PB, Challiss RA, Nahorski SR. Neuronal Ca2+ stores: activation and function. Trends Neurosci. 1996;18:299-306.
47. Dipolo R, Beauge L. Regulation of Na+-Ca2+ exchange: an overview. Ann N Y Acad Sci. 1991;639:100-111.[Medline] [Order article via Infotrieve]
48.
Haworth RA, Gokner AB. ATP dependence of calcium uptake by the Na-Ca exchanger of adult heart cells. Circ Res.. 1992;71:210-217.
49.
Shibuya I, Douglas WW. Calcium channels in rat melanotrophs are permeable to manganese, cobalt, cadmium, and lanthanum, but not to nickel: evidence provided by fluorescence changes in fura-2-loaded cells. Endocrinology. 1992;131:1936-1941.
50.
Ver Donck L, Borgers M. Myocardial protection by R 56865: a new principle based on prevention of ion channel pathology. Am J Physiol. 1991;261:H1828-H1835.
51. Wermelskirchen D, Gleitz J, Urenjak J, Wilffert B, Tegtmeier F, Peter TH. Flunarizine and R 56865 suppress veratridine-induced increase in oxygen consumption and 45Ca2+ uptake in rat cortical synaptosomes. Neuropharmacology. 1992;31:235-241.[Medline] [Order article via Infotrieve]
52. Kinskin NI, Chizhmakov IV, Tsyndrenko AY, Krishtal OA, Tegtmeier F. R56865 and flunarizine as Na+-channel blockers in isolated Purkinje neurons of rat cerebellum. Neuroscience. 1993;54:575-585.[Medline] [Order article via Infotrieve]
53. Somogyi GT, Perel JM. Biphasic effect of tricyclic antidepressants on the release of norepinephrine from the adrenergic nerves of the rabbit heart. Psychopharmacology (Berl). 1991;104:237-243.[Medline] [Order article via Infotrieve]
54.
Arita M, Wade A, Takara H, Izumi F. Inhibition of 22Na influx by tricyclic and tetracyclic antidepressants and bindings of [3H]imipramine in bovine adrenal medullary cells. J Pharmacol Exp Ther. 1987;243:342-348.
55. Cai Z, McCaslin PP. Amitriptyline, desipramine, cyproheptadine and carbamazepine, in concentrations used therapeutically, reduce kainate- and N-methyl-D-aspartate-induced intracellular Ca2+ levels in neuronal culture. Eur J Pharmacol.. 1992;219:53-57.[Medline] [Order article via Infotrieve]
56. Ogata N, Yoshii M, Narahashi T. Psychotropic drugs block voltage-gated ion channels in neuroblastoma cells. Brain Res.. 1989;476:140-144.[Medline] [Order article via Infotrieve]
57. Lavoie PA, Beauchamp G, Elie R. Tricyclic antidepressants inhibit voltage-dependent calcium channels and Na+-Ca2+ exchange in rat brain cortex synaptosomes. Can J Physiol Pharmacol.. 1990;68:1414-1418.[Medline] [Order article via Infotrieve]
58. Pandey GN, Pandey SC, Davis JM. Effect of desipramine on inositol phosphate formation and inositol phospholipids in rat brain and human platelets. Psychopharmacol Bull.. 1991;27:255-261.[Medline] [Order article via Infotrieve]
59. Mann CD, Vu MT, Hrdina PD. Protein kinase C in rat brain cortex and hippocampus: effect of repeated administration of fluoxetine and desipramine. Br J Pharmacol.. 1995;115:595-600.[Medline] [Order article via Infotrieve]
60. Schwaninger M, Schofl C, Blume R, Rossig L, Knepel W. Inhibition by antidepressant drugs of cyclic AMP response element-binding protein/cyclic AMP response element-directed gene transcription. Mol Pharmacol.. 1995;47:1112-1118.[Abstract]
61. Haass M, Richardt G, Schomig A. Potentiation of potassium-evoked noradrenaline and neuropeptide Y co-release by cardiac energy depletion: role of calcium channels and sodium-proton exchange. Naunyn Schmiedebergs Arch Pharmacol.. 1992;346:410-418.[Medline] [Order article via Infotrieve]
62.
Seyfarth M, Feng Y, Hagl S, Sebening F. Richardt G, Schomig A. Effect of myocardial ischemia in stimulation-evoked noradrenaline release: modulated neurotransmission in rat, guinea pig, and human cardiac tissue. Circ Res. 1993;73:496-502.
63. Kruger C, Haunstetter A, Gerber S, Serf C, Kaufmann A, Kubler W, Haass M. Nicotine-induced exocytotic norepinephrine release in guinea-pig heart, human atrium and bovine adrenal chromaffin cells: modulation by single components of ischaemia. J Moll Cell Cardiol.. 1995;27:1491-1506.[Medline] [Order article via Infotrieve]
64.
Dart AM, Schomig A, Dietz R, Mayer E, Kubler W. Release of endogenous catecholamines in the ischemic myocardium of the rat, B: effect of sympathetic nerve stimulation. Circ Res.. 1984;55:702-706.
65.
Miyazaki T, Zipes DP. Presynaptic modulation of efferent sympathetic and vagal neurotransmission in the canine heart by hypoxia, high K+, low pH, and adenosine. Circ Res.. 1990;66:289-301.
66.
Wakade AR, Przywara DA, Bhave SV, Mashalkar V, Wakade TD. Cardiac cells control transmitter release and calcium homeostasis in sympathetic neurons cultured from embryonic chick. J Physiol (Lond).. 1995;488:587-600.
67. Cragoe EJ Jr, Woltersdorf O Jr, Bicking JB, Kwong SF, Jones JM. Pyrazine diuretics, II: N-amidino-3-amino-5-substituted-6-halopyrazine-carboxamides. J Med Chem.. 1967;10:66-75.[Medline] [Order article via Infotrieve]
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