Synergistic Actions of Glucagon and Miniglucagon on Ca2+ Mobilization in Cardiac Cells
Abstract It has been recently shown that the physiological processing of glucagon into its C-terminal (19-29) fragment, miniglucagon, by cardiac cells was essential for the contractile positive inotropic effect of the hormone. However, the mechanisms underlying the effects of miniglucagon remained undetermined. In the present study, we assessed the effects of miniglucagon on Ca2+ homeostasis in embryonic chick ventricular myocytes. In quiescent cells, short-term applications of 0.1 nmol/L miniglucagon markedly increased the accumulation of 45Ca into intracellular compartments resistant to digitonin lysis and sensitive to caffeine. Ca2+ accumulation into the sarcoplasmic reticular (SR) store was further attested by fura 2 imaging studies on quiescent or prestimulated cells: miniglucagon potentiated Ca2+ release from the SR compartment triggered by caffeine and evoked a rise in cytosolic Ca2+ when applied on cells pretreated with 1 μmol/L thapsigargin, a specific inhibitor of the SR Ca2+ pump. Glucagon alone produced a small cytosolic Ca2+ signal that was considerably amplified by miniglucagon. The action of glucagon was mimicked by 8-bromo-cAMP and was blocked by isradipine, suggesting that it relied on the activation of L-type Ca2+ channels, via phosphorylation. We conclude that the combined actions of miniglucagon and glucagon on Ca2+ accumulation into SR stores and Ca2+ release from the same stores are likely to support the positive inotropic effect elicited in vivo by glucagon on heart contraction.
It is known that intravenous administration of glucagon, a 29–amino acid peptide derived from the pancreatic posttranslational processing of proglucagon, exerts a potent positive inotropic action on heart contraction.1 Accordingly, glucagon is prescribed in cardiac emergency in the face of β-adrenergic blockade,2 3 Ca2+ channel blocker–induced myocardial dysfunction,4 or tricyclic antidepressant poisoning.5
In a previous study, we have suggested that the positive cardiac inotropic effect of glucagon was not only due to the action of the glucagon peptide (1-29) but also relied on that of its C-terminal (19-29) fragment, referred to as miniglucagon.6 Miniglucagon is not a circulating peptide. It is generated in the extracellular medium upon incubation of heart or liver cells with glucagon.6 7 In the heart cell environment, the accumulation of miniglucagon can reach 6% of the initial glucagon concentration after 8 minutes.6 The endopeptidase responsible for the cleavage of the dibasic doublet Arg17-Arg18 in the glucagon molecule has been identified as a membranous enzyme sensitive to thiol reagents7 and has been purified from the hepatic tissue.8
We have shown that glucagon alone, under minimal degradation conditions, has no effect on the contraction of beating chick embryo ventricular cells. In contrast, a 45% increase in the amplitude of cell contractility was seen with the combination of nanomolar concentrations of glucagon and miniglucagon. Interestingly, 8-bromo-cAMP could substitute for glucagon.6
These observations implied a dual mechanism for glucagon action: one component related to the action of the glucagon peptide (1-29), involving the activation of the cAMP pathway, through either adenylyl cyclase activation or phosphodiesterase inhibition,9 10 11 and another component linked to the action of the metabolite (19-29). The latter did not rely on either cAMP or cGMP6 and remained to be defined.
Since Ca2+ plays a primary role in cardiac contraction and because we had previously shown that in liver, miniglucagon exerts a biphasic regulation on the plasma membrane Ca2+ pump ensuring Ca2+ extrusion out of the cell,12 13 14 we decided to look for a possible effect of miniglucagon on Ca2+ metabolism in heart cells. We show that miniglucagon produces a time-dependent Ca2+ accumulation into caffeine-sensitive SR compartment(s) and considerably amplifies the cytosolic Ca2+ signal triggered by glucagon.
Materials and Methods
Miniglucagon was obtained from Peninsula Laboratories Inc. Thapsigargin, caffeine, oligomycin, FCCP, penicillin-streptomycin solution, trypsin, nucleotides, and bovine serum albumin were purchased from Sigma Chemical Co. Fura 2, fura 2-AM, and pluronic acid F-127 were from Molecular Probes. Fetal calf serum was from GIBCO-BRL. PBS 2040 and medium 199 were obtained from Eurobio. Digitonin (Merck) had been recrystallized and stored as an 8 mg/mL solution in dimethyl sulfoxide. [45Ca] (5 to 30 mCi/mg) and l-[4,5-3H]leucine (40 to 60 Ci/mmol) were from ICN. Isradipine was from Research Biochemical International. Japan paper was obtained from Sennelier. An IP3 radioreceptor assay kit was purchased from Dupont-NEN.
Isolation of Chick Embryo Ventricular Cells
Fertile eggs were obtained from the Haas farm (Kaltenhouse, France). Primary monolayer cultured heart cells were prepared from 13-day-old chick embryo ventricles as previously described.6 15 The cells were dissociated by repeated cycles of trypsinization in sterile PBS 2040 medium containing 0.008% (wt/vol) trypsin. Dispersed cells were collected and diluted in ice-cold buffer A (medium 199 containing 0.1% [wt/vol] NaHCO3, 0.01% [wt/vol] l-glutamine, and 0.1% penicillin-streptomycin antibiotic solution) supplemented with 20% (vol/vol) fetal calf serum to inactivate trypsin. The cells were then filtered through sterile japan paper and centrifuged at 150g for 10 minutes to wash out the trypsin. The supernatant was discarded, and the cells were suspended in buffer A and incubated for 1 hour in humidified 5% CO2/95% air at 37°C in plastic dishes for adhesion of fibroblastic cells. Cells that remained in suspension were filtered again, and the cell suspension was diluted to 5 to 7×105 cells per milliliter in buffer A. Cells in suspension were kept in buffer A, previously bubbled with 5% CO2/95% air, at 4°C until used, up to 5 days.
Measurements of 45Ca Accumulation Into Intracellular Compartments
Isolated myocytes (5×105 cells per milliliter), suspended in buffer A supplemented with 5% (vol/vol) fetal calf serum, were plated on glass coverslips in multiwell plates and kept at 37°C in humidified 5% CO2/95% air for 24 hours. As described by Marsh et al,16 before the experiment, the cells on glass coverslips were further incubated for 24 hours in buffer A containing 120 mg/L l-[4,5-3H(N)]leucine (1 μCi/mL) in order to label cell proteins. After washing in 2×1 mL saline buffer B (mmol/L: glucose 10, NaCl 130, KCl 5, HEPES 10 [buffered at pH 7.4 with Tris base], MgCl2 1, CaCl2 2), at time zero of the experiment, the cells were immersed in 1 mL of saline buffer B containing 2 mmol/L [45Ca] (5 μCi/mL) in the presence or in the absence of 0.1 nmol/L miniglucagon and with or without 10 mmol/L caffeine. After various periods of incubation, the cells were washed two times for 10 seconds in 10 mL Ca2+-free saline buffer B at 25°C. In order to specifically evaluate the 45Ca accumulation into the intracellular stores, the cells were next subjected to digitonin lysis, which selectively disrupts the SL membranes. This procedure has been described by Altschuld et al17 and consisted of an incubation for 45 seconds at 25°C in Ca2+-free saline buffer B supplemented with 0.1 mmol/L EGTA, 10 mmol/L MgCl2, 10 mmol/L ATP, 5 μmol/L ruthenium red, and digitonin (16 μg per milligram protein). Mg2+-ATP was added to protect against hypercontracture and ruthenium red to block Ca2+ efflux from the SR. Intracellular Ca2+ accumulation was estimated from 45Ca recovered into the digitonin-resistant structures attached on the coverslips after they were dissolved in 0.2 mol/L NaOH for 2 hours at room temperature. Determination of the ratio of [3H] counts to protein concentration and simultaneous counting of [3H] and [45Ca] permitted normalization of Ca2+ uptake data per milligram cell protein. Data are expressed as mean±SEM. The significances of differences from the control values were analyzed by Student’s t test.
Fura 2 Loading and Ca2+ Imaging
The cells were plated in plastic dishes, the bottom of which was replaced by a glass coverslip that had been coated with laminin (1 μg/mL), and were incubated at 37°C in humidified 5% CO2/95% air for 17 to 24 hours. Cells, attached to laminin, were bathed in 2 mL of saline buffer B and were incubated for 20 minutes at 25°C with 1.5 μmol/L fura 2-AM (3 μL of 1 mmol/L fura 2-AM in dimethyl sulfoxide), in the presence of 1 mg/mL bovine serum albumin and 0.05% (w/vol) pluronic acid F-127 to improve fura 2 dispersion and facilitate cell loading. Cells were then washed with saline buffer B (twice, 2 mL) and allowed to incubate in the same buffer for 15 minutes at 25°C to facilitate hydrolysis of intracellular fura 2-AM. The concentration of fura 2 accumulated in the cells was estimated as described previously by Donnadieu et al.18 This consisted in comparing the fluorescence intensity of the cells loaded with fura 2-AM with that of optical oil droplets, similar in size to embryonic chick ventricular cells (0.9 to 1 μm) and loaded with known concentrations of fura 2, under the same experimental conditions (same camera gain setting, same attenuation filter). Under usual loading conditions, the average intracellular concentration of fura 2 was 15 μmol/L.
Ca2+ imaging, developed by A. Trautman in collaboration with the IMSTAR Co, was essentially described by Donnadieu et al.19 A Nikon Diaphot inverted microscope with epifluorescence was used. The light from a 100-W xenon lamp was filtered alternatively through either 350/380-nm or 360/380-nm filters to determine either [Ca2+]i or the fluorescence ratio at 360 to 380 nm (F360/F380), respectively. Fura 2 fluorescence (Nikon UV-Fluor ×40 objective) was filtered at 510 nm and recorded by an intensified CCD Photonic Science camera.
All imaging studies were performed on cells in which no spontaneous rise in [Ca2+]i was observed before experimental manipulation. We used the term “quiescent” cells as opposed to “electrically stimulated” cells.
[Ca2+]i determination was performed in quiescent cells perfused with saline buffer B, containing 2 mmol/L CaCl2 and drugs and peptides as indicated. Each fluorescence image at 350 and 380 nm was the average of four images, in order to improve the signal-to-noise ratio, and one Ca2+ ratio image was recorded every 1.2 to 3 seconds. The [Ca2+]i was calculated according to the formula of Grynkiewicz et al20 :
where Kd is the dissociation constant of the fura 2–Ca2+ complex; β is the ratio of the fluorescence of free and Ca2+-bound fura 2 measured at 380 nm and is related to the optical characteristics of the system; R is the ratio of the fluorescence signals measured at 350 nm (from which the background fluorescence at 350 nm is subtracted in the off-line analysis) and 380 nm (minus the background at 380 nm); and Rmin and Rmax are the limiting values of R in the presence of zero and saturating [Ca2+], respectively. For in vitro calibration, the value of R was determined in 14 different calibrating solutions (made in saline buffer B with various EGTA concentrations in order to vary free Ca2+ from 0.01 μmol/L to 2 mmol/L) in the presence of 15 μmol/L fura 2. Free Ca2+ concentrations of Ca2+/EGTA reference solutions were calculated by using the equiv program (Thierry Capiod, INSERM Unité 274, Orsay). An excellent linear fit of the data was obtained with Equation 1 for free Ca2+ between 0.01 and 1.5 μmol/L, which allowed, without ambiguity, the determination of Kdβ. Depending on experimental settings, the values that we determined were as follows: Rmin=0.3 to 0.4, Rmax=8 to 13.5, and Kdβ=1000 to 1700 nmol/L.
In vivo determinations of Rmin and Rmax were performed according to the protocol described by Pucéat et al.21 Maximal fluorescence was determined in the presence of 2 μmol/L 4-bromo-A23187, in cells that were previously bathed at 25°C for 1 hour in saline buffer solution containing 2 mmol/L CaCl2, without glucose and with 3 mmol/L amytal and 5 μmol/L FCCP, in order to deplete cellular ATP and to avoid cell contractures or bursts. Minimal fluorescence was obtained by bathing cells for 1 hour in saline buffer devoid of CaCl2 and containing 3.3 mmol/L EGTA with 2 μmol/L 4-bromo-A23187. Rmax and Rmin values were determined after subtraction of cell autofluorescence, measured in the presence of 1 mmol/L MnCl2. Several studies have previously reported marked differences between the in vitro and in vivo calibration data.18 In the present study, the values determined in vivo for Rmin (0.30 to 0.40) were equivalent to the in vitro values. In contrast, the Rmax values determined in vivo approached 6 and thus differed somehow from the in vitro data. This is not surprising, since intracellular systems in living cells prevent high [Ca2+]i rises. Nevertheless, in vivo calibrations gave a ratio of Rmax to Rmin of ≈20, which is an index of the dynamic range of our system that is comparable to the ranges usually reported in imaging systems, which vary from 12 to 25 (see Reference 19). Practically, [Ca2+]i calculations always took into account in vivo Rmin and Rmax and in vitro Kdβ.
In a series of experiments, cells were stimulated, and data were presented as the fluorescence ratio. Field electrical stimulation (square waves, 10-millisecond duration, amplitude 20% above threshold, 0.5 Hz) was supplied through a pair of platinum electrodes connected to the output of a HAMEG stimulator. Cells were perfused with saline buffer B containing 1.27 mmol/L CaCl2 and stimulated until a steady state level of the intracellular Ca2+ transients was achieved, before each protocol, as previously described by Bassani et al.22 Drugs and peptides were added a few seconds after interruption of stimulation. The time resolution of the imaging system was increased by restricting the microscopic field with the aid of an adjustable window and by recording only fluorescence images at 380 nm every 0.3 second (average of two images). The fluorescence at 360 nm was recorded at time zero and at the end of the experiment to check for photobleaching. Data are presented as the fluorescence ratio F360/F380, calculated after subtracting respective backgrounds.
In all experiments, applications of the different compounds were performed by including them in the perfusion medium. All tracings of [Ca2+]i or the fluorescence ratio are representative of at least 10 cells and were performed on at least two different cell isolations.
Miniglucagon Increases 45Ca Accumulation Into Caffeine-Sensitive SR Stores
Quiescent embryonic chick heart cells were incubated for various periods of time in a medium containing 2 mmol/L [45Ca], in the absence or in the presence of 0.1 nmol/L miniglucagon. This concentration was chosen since it evoked a maximal effect of the peptide on heart cell contraction.6 At the end of the incubation period, myocytes were first washed with a Ca2+-free buffer and then subjected to digitonin lysis, which selectively disrupts the SL.17 This allowed for the specific quantification of 45Ca content of digitonin-resistant structures, after elimination of the cell medium. As shown in Fig 1⇓, in the absence of miniglucagon, digitonin-resistant myocyte structures were able to accumulate an appreciable amount of [45Ca], which reached a steady state level within 1 minute. Exposing cells to miniglucagon resulted in a marked increase in the 45Ca content of internal stores, which, after a 3- or 5-minute incubation, amounted to 142% and 180% over the control level, respectively. To determine whether miniglucagon action relied on the SR stores, we examined the effect of caffeine, which has previously been shown to increase the opening probability of the Ca2+-release channels in the SR compartment.23 As shown in Fig 1⇓, after a 5-minute incubation, caffeine alone elicited a 40% decrease compared with the control level in the 45Ca content of intracellular stores. It may be noted that this gave an estimation of the caffeine-sensitive SR Ca2+ compartment versus the other intracellular Ca2+ pools, including mitochondria and nucleus. Added together with miniglucagon, caffeine completely prevented the accumulation of 45Ca into intracellular stores elicited by the peptide. These results suggest that upon miniglucagon action, Ca2+ is accumulated into cardiomyocytes being sequestered in a caffeine-sensitive SR compartment.
Miniglucagon Potentiates Caffeine-Induced Ca2+ Mobilization
The effect of miniglucagon on intracellular Ca2+ transients associated with caffeine contractures was examined in fura 2–loaded cells. Fig 2⇓ shows intracellular Ca2+ transients during electrical stimulation and caffeine contractures obtained under steady state conditions, ie, within a few seconds after interruption of electrical stimulation, according to the protocol described by Bassani et al22 (also see “Materials and Methods”). As expected,22 the application of 10 mmol/L caffeine produced a unique intracellular Ca2+ transient, larger than those observed during electrical stimulation. Miniglucagon alone had no effect (Fig 5B⇓). In contrast, the application of 10 mmol/L caffeine together with 0.1 nmol/L miniglucagon resulted in a train of intracellular Ca2+ transients (Fig 2B⇓): a mean of 4±1 transients (n=36) was observed over a period of 40 seconds. It should also be noted that when normalized as a percentage of the control amplitude, defined as the mean amplitude of the intracellular Ca2+ transients during electrical stimulation, the amplitude of the intracellular Ca2+ transients after the combined application of caffeine and miniglucagon was higher than that of the single intracellular Ca2+ transient observed with caffeine alone (127±6% [n=36] and 92±6% [n=31] of the control amplitude, respectively). The L-type Ca2+ channel blocker isradipine24 at 100 nmol/L, a concentration at which it totally abolished the intracellular Ca2+ transient during electrical stimulation (not shown), had no effect on the Ca2+ responses triggered by caffeine applied either alone or in combination with miniglucagon (Fig 2C⇓ and 2D⇓). In the presence of isradipine, the application of 10 mmol/L caffeine alone produced a unique intracellular Ca2+ transient over a period of 40 seconds, with a mean amplitude of 96±7% (n=13) compared with the control amplitude (Fig 2C⇓). Under the same conditions, the addition of miniglucagon together with caffeine triggered a mean of 4±1 intracellular Ca2+ transients (n=18) over a period of 40 seconds, with a mean amplitude of 111±3% (n=18) compared with the control amplitude (Fig 2D⇓).
Miniglucagon had no effect on IP3 production in quiescent embryonic chick heart cells, in conditions in which 100 μmol/L acetylcholine evoked a threefold increase in the IP3 level (from 20±6 to 68±22 pmol IP3 per milligram protein after a 3-minute incubation time).
Taken together, these data suggest that miniglucagon action, which leads to a higher Ca2+ loading of caffeine-sensitive SR stores, does not rely either on Ca2+ influx through L-type Ca2+ channels or on IP3-mediated Ca2+ mobilization.
We next examined the effect of miniglucagon in fura 2–loaded cells in which the SR Ca2+ storage capacity was reduced. This was achieved by pretreatment of quiescent cells with 1 μmol/L thapsigargin, a specific inhibitor of the SR Ca2+ pump.25 Preincubation with thapsigargin for 30 minutes resulted in a significant increase in basal [Ca2+]i (Fig 3A⇓) (from 45±5 to 60±5 nmol/L [Ca2+]i) and abolished completely the caffeine-induced Ca2+ signals (data not shown; see Reference 25), indicating SR Ca2+ depletion. The subsequent addition of miniglucagon caused a gradual increase in [Ca2+]i (Fig 3A⇓), which was not detected in the absence of thapsigargin (Fig 3B⇓). This observation demonstrates that miniglucagon produces an accumulation of Ca2+ in the cell that is normally immediately compensated by its sequestration into the SR compartment via the SR Ca2+ pump. It should be noted that the increase in [Ca2+]i elicited by miniglucagon in the presence of thapsigargin lasted for a few minutes only. In fact, substitution of NaCl in the extracellular medium by equimolar (130 mmol/L) LiCl prolonged the increase in [Ca2+]i that was due to miniglucagon (Fig 3A⇓), indicating that the return of [Ca2+]i to the basal level was dependent on the Na+-Ca2+ exchanger activity.
Miniglucagon Potentiates Glucagon-Induced Ca2+ Mobilization
Since, physiologically, one may expect that miniglucagon and glucagon act in concert, it was of interest to evaluate the combined effect of both peptides on [Ca2+]i.
The first series of experiments was performed in conditions in which 45Ca accumulation was observed upon miniglucagon action, ie, in cells maintained in resting conditions. The pattern of Ca2+ mobilization elicited by 30 nmol/L glucagon consisted of sporadic Ca2+ spikes (Fig 4A⇓). This small mobilization of Ca2+ produced by glucagon was considerably potentiated by 0.1 nmol/L miniglucagon (Fig 4A⇓). Note that miniglucagon alone had no effect (Fig 3B⇑).
Application of glucagon to heart cells leads to an increase in cAMP; thus, we examined the effect of the permeant analogue 8-bromo-cAMP. The nucleotide, applied alone, reproduced the effect of glucagon, eliciting single spikes of [Ca2+]i (Fig 4B⇑). Furthermore, the combination of 8-bromo-cAMP with miniglucagon reproduced the train of Ca2+ transients promoted by glucagon plus miniglucagon (Fig 4B⇑).
Since SR Ca2+ loading in quiescent cells is difficult to ascertain, a second series of experiments was performed by using the same protocol as previously described for caffeine, in which cells were prestimulated in order to ensure maximal loading of Ca2+ into the SR before application of glucagon, miniglucagon, or glucagon plus miniglucagon. Glucagon (30 nmol/L), perfused alone immediately after the interruption of electrical stimulation, produced a single Ca2+ transient over a 180-second period, the amplitude of which was 103±3% (n=29) of the control amplitude (Fig 5A⇓). In the same conditions, miniglucagon alone, at 0.1 nmol/L, did not trigger any intracellular Ca2+ signal (Fig 5B⇓). In contrast, the combination of 30 nmol/L glucagon with 0.1 nmol/L miniglucagon elicited a train of Ca2+ spikes (23±2 Ca2+ transients over a 180-second period, n=25) typical of CICR phenomenon, with a mean amplitude that was 140±5% of the control amplitude (Fig 5C⇓). These observations confirmed those made on quiescent cells.
We have previously shown that in heart cells, phosphorylation of L-type Ca2+ channels occurs as a direct consequence of cAMP production by glucagon,9 leading to a Ca2+ influx that triggers Ca2+ release from the SR compartment.26 Thus, to verify whether intracellular Ca2+ transients triggered by glucagon relied on Ca2+ influx via L-type Ca2+ channel activation, experiments were performed in which glucagon was perfused in the presence of isradipine. As shown in Fig 6A⇓, when isradipine was perfused with glucagon, after interruption of electrical stimulation, no intracellular Ca2+ transient was detected. Isradipine also totally abolished the response elicited by the combination of glucagon plus miniglucagon (Fig 6B⇓ compared with Fig 5C⇑). This result suggested that L-type Ca2+ channels are activated by glucagon, which may trigger the mobilization of intracellular Ca2+, presumably from the SR store.
This report provides further insight into the mechanism of action of glucagon and of its metabolite, miniglucagon, in the heart. Combining 45Ca uptake experiments with fura 2 imaging studies, we demonstrate a combined action of both peptides on cardiomyocyte Ca2+ homeostasis, which relies on (1) miniglucagon-induced Ca2+ loading of caffeine-sensitive SR Ca2+ stores and (2) the release of Ca2+ from these same stores, triggered by glucagon.
In quiescent cells, miniglucagon increased 45Ca content into caffeine-sensitive stores (Fig 1⇑). It is noteworthy that under our experimental conditions, an increase in 45Ca content may represent either 45Ca accumulation or increased 45Ca exchange. In the present experiments, caffeine completely suppressed the miniglucagon-induced increase in cell 45Ca loading. Since caffeine is known to trigger Ca2+ release from the SR compartment without affecting its uptake capacity, these results support the conclusion that miniglucagon produces 45Ca accumulation into the SR compartment rather than accelerating its exchange.
In fura 2–loaded cells, within a few seconds after interruption of electrical stimulation, miniglucagon potentiated [Ca2+]i mobilization induced upon caffeine contracture (Fig 3A⇑). The absence of the intracellular Ca2+ transient upon application of the peptide on cells that were responsive to electrical stimulation indicated the inability of miniglucagon to promote, by itself, the release of Ca2+ from the SR and/or to evoke action potentials by depolarizing the membrane (Fig 5B⇑). This supports the above conclusion that miniglucagon action resulted in Ca2+ accumulation into the SR stores (Figs 1⇑ and 3A⇑). Ca2+ accumulation into the SR stores evoked by miniglucagon occurred without a detectable modification of the average [Ca2+]i (Fig 3B⇑). In contrast, in the presence of thapsigargin, ie, in conditions in which the SR Ca2+ pump was inhibited, miniglucagon action resulted in a large increase in [Ca2+]i (Fig 3A⇑). These observations indicated that under normal conditions, the SR Ca2+ pump could prevent Ca2+ accumulation in the cytosolic compartment. The buffering capacity of the cardiac SR in face of trans-SL Ca2+ fluxes has been previously reported by Janczewski and Lakatta,25 who demonstrated the rapid sequestration by the SR of at least 50% of the Ca2+ entering the cells during a single postrest stimulation of guinea pig ventricular myocytes.
It may be noted that in imaging studies, miniglucagon action seemed to be immediate (Figs 2⇑, 4⇑, and 5⇑). In contrast, miniglucagon-induced 45Ca accumulation could be detected only after 3 minutes of incubation (Fig 1⇑). We interpret such differences as being due to a lower sensitivity of the isotopic technique.
On the basis of our previous reports, inhibition of the SL Ca2+ pump by miniglucagon could be responsible for Ca2+ accumulation in the cell.12 13 14 This hypothesis cannot be proved because of the lack of specific inhibitors of the SL Ca2+ pump; however, the long-term impairment of the SL Ca2+ pump, such as in diabetes,27 in genetically linked cardiomyopathy,28 or in septic shock,29 has been implicated in the development of Ca2+ overload leading to myocardial dysfunction. Alternatively, the rise in intracellular Ca2+ triggered by miniglucagon could occur via a Ca2+ influx across the SL. However, the lack of effect of isradipine on the potentiation by miniglucagon of caffeine action (Fig 2⇑) supports the hypothesis that the action of miniglucagon does not occur through L-type Ca2+ channels.
Glucagon, applied alone, induced sporadic Ca2+ transients, which were reproduced with 8-bromo-cAMP. Isradipine blocked the transients elicited in the presence of glucagon alone or glucagon plus miniglucagon. In a previous study, we showed that glucagon acted via a cAMP-dependent phosphorylation of the isradipine-sensitive L-type Ca2+ channels in cardiac cells.9 10 In addition, it has been reported that phosphorylation by the cAMP-dependent kinase of the cardiac SR CICR channel (or ryanodine receptor) leads to a more open state of the channel.30 Taken together, these observations support the proposal that glucagon, via cAMP, elicits Ca2+ influx through L-type Ca2+ channels, triggering CICR, which is also facilitated by the simultaneous phosphorylation of the CICR channel but which relies on the filling state of the SR compartment.31
Our previous data demonstrated that activation of the cAMP pathway by glucagon was necessary but not sufficient to elicit contraction.6 Accordingly, whereas it is well established that β-adrenergic agonists stimulate adenylyl cyclase activity, several reports have nevertheless demonstrated that there may be β-adrenergic–mediated pathways for increasing myocardial inotropy independent of cAMP formation.32 33 Furthermore, stimulation of other receptor types in heart cells, such as prostaglandin E1 receptors, which also induces an increase in cAMP, has no effect on contraction. Thus, the existence of cAMP pools not linked to Ca2+ and contractile regulation has been suggested.34 The present data show that the action of glucagon is dependent on both the cAMP pathway and a cAMP-independent Ca2+ loading of the SR. It is noteworthy that glucagon through this dual mechanism of action is proving more efficient than β-agonists in reversing profound myocardial depressions, in particular those induced by Ca2+ channel blocker toxicity.4
In conclusion, under physiological conditions, the positive inotropic effect of glucagon in the heart may be ascribed to the combined and distinct actions of glucagon itself, the “mother” molecule, and of miniglucagon, the “daughter” metabolite. These actions may be summarized as the ability of miniglucagon to accumulate Ca2+ into SR stores and that of glucagon to induce CICR from these stores. Preliminary experiments performed on electrically stimulated myocytes confirmed the synergistic efficiency of both peptides. Thus, continuous perfusion with 30 nmol/L glucagon alone evoked a limited increase in the amplitude of electrically stimulated intracellular Ca2+ transients (118±2% of control amplitude, n=41). In contrast, when cells were perfused with glucagon (30 nmol/L) plus miniglucagon (0.1 nmol/L), a marked rise in the amplitude of intracellular Ca2+ transients (197±8% of the control amplitude, n=52) was observed.
Selected Abbreviations and Acronyms
|β||=||fluorescence ratio of free to Ca2+-bound fura 2|
|CICR||=||Ca2+-induced Ca2+ release|
|FCCP||=||carbonyl cyanide p-(trifluoromethoxy)phenyl-hydrazone|
|Kd||=||dissociation constant of the fura 2–Ca2+ complex|
|Rmin and Rmax||=||limiting values of the fluorescence ratio in zero (min) and saturating (max) [Ca2+]|
|SR||=||sarcoplasmic reticulum, sarcoplasmic reticular|
This study was supported by the Institut National de la Santé et de la Recherche Médicale, the French Ministère de la Recherche et de la Technologie, and the Unité de Formation et de Recherche de Médecine, Créteil, Paris-Val de Marne. This material is also based on work supported by the North Atlantic Treaty Organization under a grant awarded in 1994. We thank M. Aggerbeck, R. Barouki, B. Crozatier, and R. Ventura-Clapier for helpful discussion and J. Hanoune for his permanent support.
- Received February 21, 1995.
- Accepted October 2, 1995.
- © 1996 American Heart Association, Inc.
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