The present study expands the contemporary view of mitochondria as important participants in cellular Ca2+ dynamics and provides evidence that mitochondria regulate the supply of release-competent secretory granules. Using pharmacological probes to inhibit mitochondrial Ca2+ import, the ability of mitochondria to modulate secretory activity in single, patch-clamped bovine chromaffin cells was examined by simultaneously monitoring rapid changes in membrane surface area (ΔC m) and cytosolic Ca2+ levels ([Ca2+]c). Repetitive step depolarizations or action potential waveforms were found to raise the [Ca2+]c of chromaffin cells into the 1 μm to tens of micromolar range. Inhibiting mitochondria by treatment with carbonyl cyanidep-(trifuoro-methoxy)phenylhydrazone, antimycin–oligomycin, or ruthenium red revealed that mitochondria are a prominent component for the clearance of Ca2+ that entered via voltage-activated Ca2+ channels. Disruption of cellular Ca2+ homeostasis by poisoning mitochondria enhanced the secretory responsiveness of chromaffin cells by increasing the amplitude of the transient rise and the time course of recovery to baseline of the evoked Δ[Ca2+]c. The enhancement of the secretory response was represented by significant deviation of the Ca2+–exocytosis relationship from a standard relationship that equates Ca2+ influx and ΔC m. Thus, mitochondria would play a critical role in the control of secretory activity in chromaffin cells that undergo tonic or repetitive depolarizing activity, likely by limiting the Ca2+-dependent activation of specific proteins that recruit or prime secretory granules for exocytosis.
The immediate exocytotic release of neurotransmitters from synaptic vesicles at the active zone of a synaptic bouton is governed by Ca2+ influx and the rapid collapse of microdomains of high [Ca2+]c by diffusion (Neher, 1998). Colocalization of secretory granules and Ca2+ entry sites has also been proposed for adult bovine and calf chromaffin cells (Robinson et al., 1995;Elhamdani et al., 1998). In contrast, the exocytotic release of catecholamines from secretory granules is sensitive to changes in the exogenous Ca2+ buffering capacity. This observation may be explained by indications that only a small subset of granules colocalize with Ca2+ channels (Horrigan and Bookman, 1994; Klingauf and Neher, 1997). Although there is currently little direct evidence for mitochondrial Ca2+ dynamics regulating secretory responsiveness, the concept is reasonable because mitochondria have been postulated to function as the predominant Ca2+ clearance mechanism during prolonged or repetitive stimulus-activated Ca2+influx in sympathetic neurons (Thayer and Miller, 1990; Friel and Tsien, 1994; Werth and Thayer, 1994), adrenal chromaffin cells (Herrington et al., 1996; Park et al., 1996; Babcock et al., 1997; Xu et al., 1997), gonadotropes (Hehl et al., 1996), and neuroendocrine nerve endings (Stuenkel, 1994; Giovannucci and Stuenkel, 1997).
Mitochondria can sequester large amounts of calcium and function as a cytosolic Ca2+ buffer of low affinity and high capacity (Lehninger et al., 1967; Blaustein et al., 1977;Blaustein et al., 1978; Carafoli and Crompton, 1978; Carafoli, 1979;Nicholls and Akerman, 1982; Gunter et al., 1994). Pharmacologically induced or pathophysiologically mediated mitochondrial dysfunction leads to altered Ca2+ homeostasis in neurons (Thayer and Wang, 1995; Budd and Nicholls, 1996b;Schinder et al., 1996; Wang and Thayer, 1996; White and Reynolds, 1997;Nicholls and Budd, 1998). In addition, the notion that mitochondria participate in shaping changes in cytosolic calcium concentration ([Ca2+]c) during normal cellular functioning has recently been bolstered through the simultaneous monitoring of changes in [Ca2+]c and mitochondrial free Ca2+ levels ([Ca2+]m) (Sheu and Jou, 1994; Hajnoczky et al., 1995; Sparagna et al., 1995; Jou et al., 1996; Robb-Gaspers et al., 1998; Simpson and Russell, 1998). Despite the evidence, there remains a long-standing controversy as to the functional relevance of mitochondrial Ca2+ import during neuronal activity.
Modulation of the amplitude and kinetics of the evoked Δ[Ca2+]c exerts a regulatory influence on multiple steps that control the release of neurotransmitters (Herrington et al., 1996). For example, modest increases in [Ca2+]c augment the recruitment and passage of granules through the secretory pathway via the interaction of Ca2+ ions with distinct protein targets (Bittner and Holz, 1992; von Ruden and Neher, 1993; Neher and Zucker, 1993; Zucker, 1996; Bennett, 1997; Neher, 1998). In addition, it is generally thought that the efficient secretion of neuropeptide or catecholamine requires a level of stimulatory activity that is strong enough to evoke mitochondrial participation (Peng and Zucker, 1993; Nowycky et al., 1998). In the current study, the hypothesis that mitochondria regulate secretory activity by limiting rises in [Ca2+]c and the subsequent activation of specific proteins that recruit or prime secretory granules for exocytosis was tested by monitoring stimulus-evoked changes in [Ca2+]c and the secretory activity of single bovine chromaffin cells after selective pharmacological inhibition of mitochondrial Ca2+ transport.
MATERIALS AND METHODS
Preparation of bovine chromaffin cells. Primary dissociated cells from the medullas of fresh bovine adrenal glands obtained from a local commercial slaughterhouse (Murco, Plainwell, MI) were prepared by a collagenase digestion procedure (Bittner et al., 1986). Cultures were maintained in DMEM–F-12 (BioWhittaker, Walkersville, MD) containing 10% heat inactivated FCS. Cells were cultured as monolayers on collagen-coated glass coverslips (32 μg/ml in 0.01 N HCl), which formed the bottoms of 35 mm culture dishes (500,000–1,000,000 cells per dish). Before the start of an experiment, culture medium was replaced by superfusion with physiological saline for ∼20 min. Experiments were performed 1–8 d after the preparation of the cell cultures.
Electrophysiological recording ofIca and Cm .Standard whole-cell and perforated patch-clamp methods were used to evoke and record calcium currents and measure small, time-resolvable changes in membrane capacitance (ΔC m) from single chromaffin cells using a modified Axopatch 200A amplifier (Axon Instruments, Foster City, CA) and phase-tracking software (Pulse Control; Drs. Jack Herrington and Richard Bookman, University of Miami Medical School, Miami, FL). The ΔC m was monitored by applying a sine wave (60 mVp-p at 1201 Hz) to a holding potential of −90 mV. Sixteen samples per sinusoidal period were used to compute one C mpoint each 6.6 msec, and calibration pulses (100 fF and 500 kΩ) were generated at the beginning of each trace. A train of 8 or 12 50–100 msec step depolarizations from −90 to 10 mV at 0.2 or 0.5 sec intervals was applied to evoke I Ca, Δ[Ca2+]c, and ΔC m. For standard whole-cell patch recordings, pipettes were constructed out of 1.5 mm outer diameter (o.d.) capillary glass (Drummond Scientific, Broomall, PA) coated with Sylgard elastomer and fire polished to resistances of 2.5–7 MΩ. The standard intracellular recording solution contained (in mm):N-methyl-d-glucamine-Cl 128, HEPES 40, NaCl 10, Mg-ATP 4, GTP 0.2, Tris-EGTA 0.1, and fura-2, 0.15, pH adjusted to 7.1. For some experiments, 1 mm n-hydroxyethylethylenediaminetriacetic acid (HEDTA) or 10 μm ruthenium red (RR) was added to this solution. When necessary, osmolarity was maintained by ionic substitution. Conventional whole-cell recording was used for most experiments. For experiments in which cells were loaded with furaptra AM or stimulated by action potentials (see below), the perforated patch-clamp configuration was used. For these experiments, pipettes were constructed out of 1.5 mm o.d. borosilicate glass (catalog #TW150F-4; World Precision Instruments, Sarasota, FL). The pipette solution contained (in mm): cesium methanesulphonate 140, HEPES 10, MgCl2 1, EGTA 0.1, and amphotericin B 0.26, pH adjusted to 7.2 with CsOH. A concentrated stock solution of amphotericin B (30 μg/μl in methyl sulfoxide) was made fresh for each experiment and used within 1 hr. For recording of I ca, the superfusion solution was changed to a solution containing (in mm): tetraethylammonium chloride 137, CaCl2 10, MgCl2 2, HEPES 10, and glucose 19, pH adjusted to 7.15 with Tris. Test solutions containing mitochondrial inhibitors (0.5–1 μmcarbonyl cyanide p-(trifuoro-methoxy)phenylhydrazone (FCCP), 1 μm oligomycin, 10 μmantimycin and 10 μm oligomycin, or 100 μm CdCl2 were applied by local perifusion through a length of fused silica tubing (inner diameter of 300 μm) (PolyMicro Technologies, Inc., Phoenix, AZ) placed ∼50 μm from the cell. All compounds were purchased from Sigma (St. Louis, MO).
Action potential clamp. Action potentials were evoked by brief current injection or by application of the nicotinic agonist DMPP (2 μm), and membrane voltage changes were recorded in the standard whole-cell configuration under the current-clamp mode of an Axopatch 200A amplifier with a sampling rate of 10 kHz. The pipette solution contained (in mm): KCl 135, HEPES 10, glucose 10, MgCl2 2, and EGTA 0.250, and pH was adjusted to 7.2. Action potentials from four cells were digitally recorded and averaged to produce a stimulus waveform used for subsequent patch-clamp experiments and were applied as a single stimulus or in trains of 144 action potentials at 5 Hz. For these experiments, the sampling rate was adjusted to match that of the stimulus waveform (100 μsec/C m point). FifteenC m points were determined every 1.76 sec [after every 12th action potential (AP)]. The current output of the amplifier was transformed by a digital pulse code audio processor (PCM-701ES; Sony, Tokyo, Japan) and stored for playback on a video cassette recorder (Betamax SL-2700; Sony).
Epifluorescence measurement of [Ca2+]c. To determine Δ[Ca2+]c, 150 μm fura-2 or furaptra was included in the intracellular recording solution, and the fluorescence was monitored using dual wavelength microspectrofluorometry (SPEX Industries, Edison, NJ). Individual chromaffin cells were optically isolated using a 10 μm pinhole stop and then illuminated by epifluorescence through a 40× oil immersion objective (NA of 1.30) with alternating excitation wavelengths of 340 and 380 nm. The emission at 510 nm was measured by photomultiplier (15–100 msec/point), and the [Ca2+]c was obtained using the ratiometric method (Grynkiewicz et al., 1985): Fura-2 and furaptra signals were calibrated using a solution similar to the intracellular patch recording solution and containing either nominal (10 mm EGTA, no added Ca2+) or saturating (2.9 mm) free [Ca2+] and constant free [Mg2+] of 0.74 mm(determined using Patcher's Power Tools XOP; Dr. Francisco Mendez, Department of Membrane Biophysics, Max-Planck-Institute for Biophysical Chemistry, Gottingen, Germany). After subtraction of background autofluorescence measured before rupture of the cell membrane patch,R min,R max, and β were determined to be 0.35, 11.4, and 9.6 for fura-2, and 0.47, 6, and 8.3 for furaptra, respectively. A K D value for fura-2 of 224 nm was taken from the literature (Grynkiewicz et al., 1985), and β was determined by multiplyingK D by the ratioF 0/F s. In experiments in which the perforated patch-clamp configuration was used, cells were loaded by perifusion with a physiological saline solution containing 1 μm fura-2 AM or furaptra AM. In these cells, background autofluorescence was determined after attainment of whole-cell configuration and washout of the dye. The dissociation constant (K D) of furaptra has been estimated to range between 20 and 53 μm (Raju et al., 1989; Hurley et al., 1992;Naraghi, 1997; Xu et al., 1997). Under our experimental conditions, theK D of furaptra was estimated to be 20 μm by matching the Δ[Ca2+]c for a specific Ca2+ influx as determined by fura-2 to that evoked by the same influx in furaptra loaded cells, and substituting R, R min,R max, and β into the equation above to solve for K D.
Unless otherwise indicated, experiments were performed in 10 mm external [Ca2+] using conventional whole-cell patch-clamp configuration to evoke and monitor both ΔC m andI Ca. The general experimental paradigm and nomenclature used is illustrated in Figure1, A and B, in which both the cumulative change in membrane capacitance after each step depolarization (ΔC m Pn, where n indicates the position of a particular step depolarization within a pulse train) and the maximal ΔC m(ΔCm max) were determined before and after drug application. The value of the ΔC m Pn, which represents the C m change with respect to the basal C m value, was measured ∼20 msec after cessation of the step depolarization and includes any exocytotic activity that occurs during the interpulse intervals. The ΔC m maxreflects the largest value achieved within 30 sec after initiation of the stimulus train. The I Cacorresponding to each step depolarization was integrated, and the cumulative charge of entering Ca2+ ions (ΣQ Ca) was related to the ΣΔC m to investigate modulation of the Ca2+–exocytosis relationship. The bovine chromaffin cells used in the present study had a mean diameter of 15.2 μm and a resting whole-cellC m of 6.5 ± 0.6 pF (n = 36). Under conventional whole-cell patch-clamp conditions, application of an initial train of repetitive depolarizations induced an averaged cumulative, time-integrated Ca2+ influx of 161 ± 46 pC and a ΔC mmax of 248 ± 49 fF (n = 14). This increase corresponded to the exocytotic fusion of ∼65 secretory granules (3.8 fF/granule), assuming the average diameter of a single chromaffin granule is 0.356 μm with a specific membrane capacitance of 9 fF/μm2 (Albillos et al., 1997; Plattner et al., 1997). In control records, diminishment of theC m step amplitude evoked by each pulse during the train was observed in 57% of the cells. In these cells, the cumulative ΔC m evoked by the final step depolarization (ΔC m P8 or ΔC m P12) and the ΔC m maxgave comparable values. This diminishment in the amplitude of theC m steps has been postulated to reflect the activity-dependent depletion of a pool of release-ready granules or a short-term change in the Ca2+–exocytosis relationship (Horrigan and Bookman, 1994; Engisch and Nowycky, 1996; Engisch et al., 1997). The remaining cells were found to exhibit a further average increase inC m (83 ± 25 fF) that persisted for 3.4 ± 1.9 sec after termination of the stimulus train (n = 6). This persistence of secretion may represent the exocytosis of release-ready granules that require the diffusional overlap of multiple Ca2+ domains or granules that require Ca2+-dependent recruitment and/or priming steps before fusion. Both types of responses were included in the averaged data relating Ca2+ influx andC m Pn andC m maxincreases.
Because secretory response characteristics of a single chromaffin cell may change in a time- or activity-dependent manner,C m changes in response to successive pulse trains were also monitored. As shown in Figure 1 C, there was a decline in both the amplitude of the depolarizing pulse-evoked Ca2+ currents and in the ΔC m max with sustained dialysis (n = 5). Although the rate with which responsiveness declined was variable between cells, a significant enhancement of the ΔC m maxbetween successive pulse trains applied at 2 min intervals under control conditions was rarely observed.
Effect of FCCP on the stimulus-evokedCm response
To determine the contribution of mitochondrial Ca2+ buffering to the control of catecholamine release, FCCP was used to dissipate the proton gradient across the inner mitochondrial membrane and reduce the electrochemical driving force (ψm + ΔpH) for mitochondrial Ca2+ import. The [Ca2+]c andC m changes evoked by repetitive stimuli before and during treatment with 0.5–1 μm FCCP were then compared. Neither FCCP (0.5–1 μm) nor oligomycin (1–10 μm) alone had any significant effect on basal levels of [Ca2+]cand C m. However, as shown in Figure2 A, the application of FCCP was found to potentiate the ΔC m maxevoked by repetitive step depolarizations sevenfold over that of the control C m response (n= 14; p < 0.001). Although FCCP treatment potentiated the evoked secretory response in nearly all cells tested, this enhancement varied from cell to cell in both magnitude and time course. The ΔC m for Cell 3 is shown on an expanded time scale in Figure 2 B and includes the corresponding Δ[Ca2+]c and first and final I Ca evoked by the stimulus trains. This type of response was observed in 43% of cells and demonstrated a moderate or profound increase in ΔC m Pn during the pulse train, often despite decreased Ca2+ influx. By applying depolarizing stimuli to this cell at 1 Hz, it can be seen that, after FCCP treatment, the majority of the ΔC m Pnincrease is not synchronized with Ca2+entry and occurs during the interpulse intervals. Because we are unable to isolate the C m change evoked by active Ca2+ influx from that of the persistent C m rise, we have focused on comparing the cumulative C m changes evoked by Ca2+ influx (as a measure of the Ca2+–exocytosis relationship) between control and FCCP-treated cells. Despite decreased Ca2+ influx during FCCP treatment, there was little difference in the magnitude of the [Ca2+]c during the stimulus as reported by the fura-2 dye. This apparent discrepancy may be explained by an inability of the fura-2 dye to accurately report the large changes in [Ca2+]c induced by the strong stimuli used and further compounded by enhancement of the [Ca2+]c by FCCP (see next section). In the remaining cells, enhancement during the train was not readily evident and, in some cases, appeared to be diminished by FCCP treatment. However, when the ΔC m Pn was normalized to account for diminished Ca2+influx, an enhanced Ca2+–exocytosis relationship was revealed (see below). It is important to note that, in most cells treated with FCCP, the majority of the ΔC m occurred after the stimulus train had ended. This persistent rise in the ΔC m often lasted for tens of seconds after voltage-dependent Ca2+ influx. It is unlikely that these effects resulted from depletion of cellular ATP levels or rundown of the plasma membrane Ca2+ pumps. Because FCCP treatment can elicit ATP consumption by reversal of theF 0-F 1ATP synthase, use of FCCP was always coupled with 1 μm oligomycin, a specific blocker of the mitochondrial ATP synthase (Budd and Nicholls, 1996a). Both the cytosolic ATP concentration (4 mm) and the pH (40 mm HEPES, pH. 7.1) were controlled by the use of the whole-cell recording configuration.
As shown in Figure 3 A, the average ΔC m maxevoked before FCCP treatment was 248 ± 49 fF, whereas that evoked during treatment with FCCP was 1743 ± 275 fF (n = 14). Unlike the modest increases inC m max induced in control cells, the FCCP-treated cells demonstrated a considerable enhancement of the ΔC m max. The persistent exocytotic response was maximal within tens of seconds after cessation of the stimulus train. On average, the ΔC m max after FCCP treatment corresponded to the fusion of 1–2% of the estimated total granule content of a chromaffin cell (26,000–30,000 granules per cell) (Plattner et al., 1997) and an apparent increase in the number of granules available for release by this stimulus train from 65 to 457 granules. Because it is estimated that each bovine chromaffin cell contains 496 secretory granules that are either docked or in close proximity to the plasma membrane (Plattner et al., 1997), an interpretation is that the stimulus protocol induced the fusion of greater than 90% of this pool. Figure 3 A shows that, in addition to the ΔC m maxenhancement, the final ΔC m Pn(ΔC m P8 or ΔC m P12) was also enhanced compared with control, indicating that increased secretory responsiveness developed during the stimulus train. As shown in Figure 3 B, the FCCP-induced enhancement of the ΔC m max was accompanied by a significant deviation from a standard relationship equating Ca2+ influx and the stepwise ΔC m in bovine chromaffin cells (Engisch and Nowycky, 1996; Engisch et al., 1997). The enhancement of the Ca2+–exocytosis relationship developed during the stimulus train, such that the latter ΔC m Pn in the series, measured immediately after the termination of each depolarization, were enhanced significantly compared with control ΔC m Pn steps. Figure 3 B compares the Ca2+–exocytosis relationship evoked by eight step depolarizations in FCCP-treated cells with that of control cells and to a line fit to the standard Ca2+–exocytosis relationship (n = 8). This enhancement occurred despite rundown in the total amount of Ca2+ influx that accompanied FCCP treatment during the stimulus train (see below).
Additional experiments were performed to verify that the FCCP-induced enhancement of the secretory response was dependent on Ca2+ influx. As shown in Figure4 A, application of 100 μm Cd2+ blocked Ca2+ influx through voltage-dependent Ca2+ channels during FCCP treatment and abolished the ΔC m andI Ca (Fig. 4 A). A subsequent stimulus train after removal of Cd2+, but in the continued presence of FCCP, evoked an enhanced ΔC m max(2233 ± 674 vs 292 ± 123 fF; n = 3). In addition, in a limited number of cells, removal of FCCP could restore the C m response to control levels (Fig. 4 B), consistent with the notion that the FCCP-induced increases in the evoked ΔC m were mediated by reduced mitochondrial Ca2+ import rather than collapse of cellular ATP levels. However, after the robust, FCCP-enhanced secretory response and prolonged elevation of [Ca2+]c, the majority of the cells treated did not respond to a subsequent stimulus train in the continued presence of FCCP.
FCCP-induced changes in [Ca2+]c dynamics
The FCCP-induced enhancement of the secretory response was accompanied by an alteration in the magnitude and time course of the [Ca2+]c response. To further establish that the increased secretory responsiveness during FCCP treatment resulted specifically from a derangement of Ca2+ homeostasis, the effect of FCCP on the Δ[Ca2+]cevoked by repetitive step depolarizations was more thoroughly examined using the Ca2+-sensitive fluorescent probes fura-2 and furaptra. The resting level of [Ca2+]c of chromaffin cells in standard or oligomycin-containing saline monitored with fura-2 was typically 125 ± 24 nm(n = 8). As shown in Figures 2 B and5, A and B, [Ca2+]c increased in control cells to a plateau level by the third or fourth step depolarization in a train of depolarizing pulses. On average, the Δ[Ca2+]c was estimated by fura-2 to be 972 ± 228 nm and returned to a level just above that of prestimulus with an average time constant of 18 ± 3 sec after cessation of the stimulus (n = 8). After treatment with FCCP, the magnitude of the Δ[Ca2+]c was increased (1804 ± 457 nm; p< 0.017; n = 8) compared with control values, and the recovery of the [Ca2+]c was markedly slowed (t 1/2 = 81 ± 29 sec; n = 5) or remained elevated. Moreover, the majority of the increase over control values occurred after the stimulus train and was represented by a slow upward drift of the [Ca2+]c level. Unexpectedly, the Δ[Ca2+]c in FCCP-treated cells also reached a plateau during the stimulus, indicating that the decrement of the Δ[Ca2+]c during influx was not solely a function of mitochondrial uptake.
Because the estimated K D for fura-2 is 224 nm, it is expected that when [Ca2+]c rises to ∼2 μm, ∼90% of fura-2 will become saturated (Augustine and Neher, 1992; Zhou and Neher, 1993). We, therefore, suspected that the loss of proportionality between Ca2+ influx evoked by repetitive depolarizations and dye fluorescence resulted from a loss of sensitivity because of dye saturation and that fura-2 measurements may underestimate the Δ[Ca2+]c in response to repetitive depolarizations. To address this concern, chromaffin cells were loaded with furaptra, which has a 100- to 250-fold lower affinity for Ca2+-binding. After loading with furaptra AM, the average increase in [Ca2+]c evoked by repetitive step depolarizations under perforated patch-clamp conditions was estimated in control cells at 7.2 ± 1.6 μm (n = 7). FCCP-treatment, however, significantly enhanced the evoked Δ[Ca2+]c, which averaged 12.9 ± 2.5 μm (n= 3). An example of the evoked changes in [Ca2+]c for a furaptra-loaded cell with a 15 μm diameter, before and during FCCP treatment, is shown in Figure 5 C. The cytoplasmic Ca2+-binding ratio for bovine chromaffin cells is estimated to be near 100 (Neher and Zucker, 1993). Assuming that the estimated endogenous buffering capacity of a chromaffin cell does not change significantly above 2 μm, the depolarization-mediated Ca2+ influx of 580 pC for this cell should have raised the [Ca2+]c to ∼20 μm, assuming an accessible cell volume of 1450 μm3 [1767 μm3 × 0.85 (Xu et al., 1997)]. However, the [Ca2+]c increased to a value just over half that estimated (11.7 μm). After mitochondrial uncoupling by FCCP, however, a second train of depolarizations mediating a cumulative influx of 456 pC now raised the [Ca2+]c to 18.8 μm, indicating that under the control conditions mitochondria acted to rapidly buffer changes in [Ca2+]c in the micromolar range. Because experiments using furaptra were performed after loading of the indicator dye as the membrane-permeable form, no absolute estimate of the cytosolic concentration of the dye was made, making a quantitative estimate of the Ca2+-binding capacity of furaptra and of the contribution of mitochondria Ca2+buffering under control conditions equivocal. Interestingly, in four of seven cells, FCCP treatment revealed an additional increase in [Ca2+]c after cessation of the stimulus train (Fig. 5 D). This delayed rise in [Ca2+]csuggested that mitochondria may also sequester Ca2+ released from an unidentified intracellular site or, when uncoupled, may themselves release stored Ca2+ in response to Ca2+ influx through voltage-activated Ca2+ channels, perhaps via permeability transition. However, the subpopulation of furaptra-loaded cells that exhibited the delayed [Ca2+]c rise after the stimulus train was not included in the estimates of [Ca2+]c rise in response to influx.
Other inhibitors of mitochondrial Ca2+ import augment the secretory response
Under whole-cell patch-clamp configuration, enhanced secretory responsiveness, similar to that observed after uncoupling mitochondria with FCCP, could be induced by the inhibition of mitochondrial Ca2+ import using two other mitochondria-specific poisons, each with distinct mechanisms of action. The data from these experiments are summarized in Figure6. When 10 μm RR, a relatively specific blocker of the mitochondrial Ca2+ uniporter, was introduced through the patch pipette, the average evoked ΔC m max was 1325 ± 75 (n = 4). This value was significantly larger than that evoked by the same stimulus under standard control conditions (248 ± 49 fF). When FCCP was applied in combination with RR, there was observed no further enhancement of the secretory response (1100 ± 289 fF; n = 4). The lack of an additive effect of these compounds on the secretory response suggests that these probes act at the same intracellular compartment.
In addition to RR, we used a combination of 10 μmantimycin A1 and 10 μm oligomycin to reduce the inner mitochondrial membrane potential (ψm) and, hence, the electromotive force for mitochondrial Ca2+ import. Antimycin A1 is an antibiotic substance that specifically inhibits electron flow between cytochrome b and c1 of the respiratory chain and blocks proton gradient generation at site 2. The ΔC m maxevoked by repetitive step depolarizations before and during chemical hypoxia induced by 3–5 min of treatment with antimycin was 259 ± 111 and 711 ± 280 fF, respectively (n = 6;p < 0.04).
In addition to their effects on cellular Ca2+ homeostasis, inhibitors of mitochondrial function have been shown to alter the cellular levels of ATP–ADP, H+, Na+, and reactive oxygen species (ROS) in intact cells, each of which may affect the exocytotic response (Carriedo et al., 1998; Tenneti et al., 1998). Whereas the concentrations of ATP and ionic constituents can be maintained at relatively constant levels by the whole-cell patch-clamp configuration, significant amounts of ROS may be produced under our experimental conditions. To confirm the hypothesis that the increased secretory responsiveness after mitochondrial dysfunction resulted from a perturbation of [Ca2+]c dynamics, we attempted to “reconstitute” mitochondrial Ca2+ buffering capacity by including in the patch pipette solution a Ca2+ buffer with an affinity and capacity similar to that estimated for the mitochondrial component. It has been estimated that the mitochondria of bovine chromaffin cells have the capacity to sequester a total cytoplasmic Ca2+ load of 1 mm(Xu et al., 1997). Accordingly, 1 mm HEDTA was chosen to simulate the mitochondrial buffering capacity because it has an estimated buffering range of 1.3–8 μm under our experimental conditions. FCCP-induced uncoupling of mitochondrial Ca2+ import had no significant effect on either the ΔC m or the Δ[Ca2+]c when HEDTA was included in the patch pipette solution. As shown in Figure 6, the average peak values for these before and after FCCP-treatment were 738 ± 425 fF and 456 ± 186 nm, respectively (n = 4). The increased average ΔC m max in HEDTA-loaded cells resulted from an enhancement of theI Ca, and inclusion of one cell in the data set that had uncommonly large I Ca(>1 nA). Moreover, the Ca2+–exocytosis relationship between control and HEDTA-treated cells was not significantly different (data not shown). Thus, the enhanced secretory activity resulting from mitochondrial dysfunction was abolished by the intracellular application of an exogenous low-affinity Ca2+ buffer. Although the role of antioxidants was not directly tested, the above results suggest that the effect of mitochondrial inhibitors on secretory activity primarily results from a perturbation of Ca2+homeostasis rather than from ROS generation.
To determine whether mitochondrial Ca2+ import controls secretory activity under physiological conditions, experiments were performed using action potential waveforms as the depolarizing stimulus under perforated patch configuration and with the external Ca2+concentration reduced from 10 to 2.2 mm. In this manner, cytosolic proteins and the endogenous Ca2+buffer capacity of the cell are maintained and the influx driven by the depolarizing stimulus more closely approximates the physiological situation. Moreover, in rat and guinea pig preparations, splanchnic nerve or muscarinic stimulation has been shown to elicit bursts of action potentials (1–30 Hz) and the exocytotic release of catecholamine (Brandt et al., 1976; Kidokoro and Ritchie, 1980;Kajiwara et al., 1997; Inoue et al., 1998). In addition, acetylcholine-mediated depolarization of bovine chromaffin cells can induce trains of action potentials capable of inducing catecholamine secretion (Douglas et al., 1967). Accordingly, we recorded action potentials from chromaffin cells under current clamp and averaged them to use as a stimulus waveform (AP). This AP waveform was similar to those reported by others recorded from bovine and mouse chromaffin cells (Fenwick et al., 1982; Zhou and Misler, 1995; Moser, 1998). The prerecorded AP was then applied in a train at 5 Hz (144 APs) from a holding potential of −50 mV to evoke Ca2+influx, Δ[Ca2+]c, and ΔC m. An averagedI Ca activated during a single AP is shown in Figure 7 A. The AP-evoked I Ca had a peak amplitude and current integral of 450 ± 139 pA and 1.18 ± 0.41 pC (n = 6), respectively, and was completely blocked by the local application of 100 μmCd2+ (data not shown). TheI Ca activated at −16 mV reached a peak amplitude at −6 mV during the falling phase of the AP and had a half-width of 2.5 msec. The AP-evoked ΔC m did not exhibit activity-dependent depression of the ΔC m and was significantly enhanced from that predicted by the standard relationship determined using patterns of step depolarizations (see Discussion). For example, under control conditions, a train of 144 APs evoked a cumulative influx of ∼170 pC (530 × 106Ca2+ ions) and ΔC m and Δ[Ca2+]c of 673 ± 246 fF and 361 ± 67 nm(n = 6). This may indicate that, during AP-mediated secretory activity, granule recruitment is matched to support continued exocytosis or that trains of APs may activate a facilitationI Ca. The latter possibility was excluded because there was observed no facilitation of the Ca2+ currents evoked by repetitive application of APs, consistent with recent work that demonstrated theI Ca of adult bovine chromaffin cells do not facilitate (Engisch et al., 1997; Elhamdani et al., 1998). To estimate the contribution of mitochondrial Ca2+ import to the secretory response evoked by a physiological stimulation, we compared the AP-evoked ΔC m response before and during FCCP treatment. As shown in the representativeC m records in Figure 7 B and averaged data in Figure 7 C, a reduction of mitochondrial Ca2+ buffering capacity reversibly potentiated the ΔC m. The average ΔC m evoked during FCCP treatment was 1414 ± 466 fF (n = 6). After removal of FCCP, the ΔC m returned to 688 ± 324 fF, a value not significantly different from that evoked before FCCP treatment (n = 4). Although the increased responsiveness was less than that observed under stimulatory conditions that drive secretion maximally, these data indicate that mitochondrial import can contribute significantly to the control of secretory granule exocytosis during repetitive stimulatory activity.
In response to multiple step depolarizations, the [Ca2+]c of bovine chromaffin cells was found to escalate from ∼0.1 to 1–20 μm, a range of [Ca2+]c in which mitochondria dominate Ca2+ clearance (Herrington et al., 1996; Xu et al., 1997). Pharmacological suppression of this low-affinity buffering mechanism by treatment with FCCP, RR, or antimycin–oligomycin rapidly potentiated the Δ[Ca2+]c and resulted in a threefold to sevenfold increase in the pool of secretory granules releasable by a standard stimulus pattern. Treatment with FCCP and RR in combination had no additive effect on secretion, suggesting that these compounds act on the same intracellular compartment. Simulating the endogenous buffering capacity of mitochondria by introduction of a low-affinity high-capacity Ca2+ buffer blocked the FCCP-induced enhancement of secretion. These findings indicate that mitochondria play an important role in the control of secretory activity in chromaffin cells.
Patterned activity has been shown to induce short-term changes in the secretory responsiveness of bovine chromaffin cells such that the rise in C m evoked by repetitive stimulations can deviate from that predicted by a simple relationship describing Ca2+ influx and ΔC m (Engisch et al., 1997). To account for this deviation, it has been proposed that the efficacy with which Ca2+ can elicit a ΔC m may be enhanced by the activation of intracellular Ca2+ release or secretory granule mobilization, diminished by the recruitment of rapid Ca2+ clearance mechanisms (Hehl et al., 1996), or by desensitization of the secretory apparatus (Stuenkel and Nordmann, 1993; Hsu et al., 1996). In the present study, controlC m responses closely followed the standard Ca2+–exocytosis relationship described by Engisch and Nowycky (1996) and Engisch et al. (1997), but deviated from the predicted relationship when mitochondrial Ca2+ import was inhibited. The initial exocytotic steps, however, were not enhanced, suggesting that the exocytotic event per se was not directly modulated by mitochondrial inhibitors. In addition to the enhanced ΔC m that was directly coupled to Ca2+ influx, repetitive activity in the presence of mitochondrial inhibitors also produced a persistent, “asynchronous” rise in C m that followed the cessation of the stimulus, but was, nevertheless, dependent on previous Ca2+ entry. This persistent rise in C m likely resulted from global rises in [Ca2+]c and the recruitment for exocytosis of granules that are not located near the sites of Ca2+ influx.
The increased efficacy with which a train elicits secretion after FCCP treatment is represented by an increase in the number of granules recruited or available for release rather than a direct effect on the Ca2+ sensitivity of the release machinery. This interpretation is consistent with the observation that the ΔC m P1 was unaffected in both control and FCCP-treated cells, that significant enhancement of the Ca2+-exocytosis relationship developed during the stimulus train, and that most of the increase occurred during the interpulse intervals and after cessation of the stimulus train. Placed within the context of a two-step model for secretion in chromaffin cells (Heinemann et al., 1993, 1994; Smith et al., 1998), the enhanced magnitude and prolonged elevation of the [Ca2+]c associated with mitochondrial inhibition may act to drive the recruitment of granules into a releasable pool. Thus, long-term micromolar increases in [Ca2+]c evoked by strong repetitive stimuli in the presence of FCCP would act to either (1) increase the throughput of granules to the final exocytotic event in a stepwise enzymatic cascade in which granules exist in pools or states of varying releasability, or (2) increase the number of release sites that are activated by a given stimulus, in the way photolytic release of Ca2+ can evoke a massive secretory response. Implicit in the latter of these possibilities is the suggestion that global increases in [Ca2+]c must also act to drive persistent exocytosis from release sites distributed over the plasma membrane. These two scenarios are not necessarily exclusive, and experimental manipulations of mitochondrial Ca2+ uptake may provide useful tools for probing the recruitment and availability of secretory granules for exocytotic release.
A remarkable finding of the present study was that blocking mitochondrial Ca2+ import was found to enhance changes in C m and [Ca2+]c when patterns of stimulation using natural (action potential) waveforms under conditions that preserved the physiological milieu were used to evoke secretion. For example, FCCP treatment potentiated the secretory response evoked by action potentials over twofold compared with control responses. Thus, even under conditions of moderate Ca2+ influx, mitochondria play a prominent role in limiting exocytotic activity in bovine chromaffin cells, primarily by rapidly clearing from the cytosol Ca2+ that accumulates during repetitive stimulations. Engisch et al. (1997) reported that enhancement is induced under conditions of minimal Ca2+entry. This is consistent with our observations that the total ΔC m evoked by trains of APs under control conditions was enhanced more than twofold from the predicted Ca2+–exocytosis relationship and, further, did not exhibit the use-dependent depression commonly observed when square-wave depolarizations were used to evoke secretion. Thus, it appears that natural waveforms or patterns of stimulation are more efficient at eliciting exocytotic fusion (Zhou and Misler, 1995;Engisch et al., 1997). Furthermore, after inhibition of mitochondrial Ca2+ import, the evoked ΔC m was enhanced more than fourfold from the standard Ca2+–exocytosis relationship, demonstrating that mitochondria normally limit secretory activity under physiologically relevant conditions.
The enhanced secretory responsiveness may reflect the time- and activity-dependent activation of specific Ca2+-regulated proteins and their effectors whose function is to regulate the supply of release-competent secretory granules. For example, members of the protein kinase C family are one set of promising candidates for this regulatory control because they are activated by elevation of [Ca2+]c in chromaffin cells (TerBush et al., 1988), and phorbol ester treatment has been shown to induce a long-lasting enhancement of the secretory response (Bittner and Holz, 1993; Gillis et al., 1996; Billiard et al., 1997; Cox and Parsons, 1997; Misonou et al., 1998; Smith et al., 1998).
Presynaptic Ca2+ clearance by mitochondria may play a general role to regulate synaptic strength. This notion is based on long-standing information detailing the abundance of mitochondria at nerve endings (Fried and Blaustein, 1978), the multiple Ca2+ transport mechanisms associated with mitochondria (Sparagna et al., 1995; Gunter et al., 1998), and the ability to increase transmitter release when mitochondrial Ca2+ transport is inhibited (Alnaes and Rahamimoff, 1975; Melamed-Book and Rahamimoff, 1998). Recently,Peng (1998) demonstrated that mitochondria are an important, frequency-dependent mechanism for Ca2+removal after repetitive firing at peptidergic presynaptic terminals of bullfrog sympathetic ganglia. Also, a direct demonstration of activity-dependent mitochondrial Ca2+transport at the lizard neuromuscular junction was resolved by David et al. (1998). Using Oregon Green-5N and Rhod-2 dyes in combination to simultaneously monitor Δ[Ca2+]c and Δ[Ca2+]m, respectively, they found that repetitive stimulations (30–50 APs) raised the [Ca2+]mafter the onset of a rise in [Ca2+]c and demonstrated an enhancement of the Δ[Ca2+]c after interruption of Δ[Ca2+]m. Buffering of [Ca2+]c by mitochondria may also play an important role at some mammalian central synapses. For example, Borst and Sakmann (1996) estimated that, at a central fast synapse in the rat brain, ∼60 Ca2+ channel openings were required to evoke the release of neurotransmitter and demonstrated that the release event was subject to modulation by relatively slow-acting Ca2+ buffers. The sensitivity of the exocytotic response to changes in the Ca2+buffering capacity underscores the potential contribution of presynaptic Ca2+ clearance by mitochondria to modulate synaptic strength.
A key question to be addressed in future studies is whether mitochondrial Ca2+ import in neuroendocrine cells is a regulated process. Although this study has primarily focused on the effects of inhibition of Ca2+ import, Ca2+ efflux from the mitochondrion may function to produce a prolonged low-level elevation in [Ca2+] that could support recruitment and priming of fusion-competent secretory granules. For example, the use of inhibitors of mitochondrial Ca2+transport showed that, during tetanic stimulation of the crayfish neuromuscular junction, neurotransmitter release was enhanced and demonstrated that mitochondrial Ca2+efflux underlies the generation of post-tetanic potentiation (Kamiya and Zucker, 1994; Tang and Zucker, 1997). Accordingly, the notions that mitochondria normally can function to limit or sustain and augment secretory activity are not necessarily mutually exclusive concepts.
This work was supported by National Institutes of Health Grant NS36227 to E.L.S. We thank Drs. James Herrington, Ronald Holz, Mary Bittner, and Brandi Soldo for valuable discussion.
Correspondence should be addressed to David Giovannucci's present address: Department of Pharmacology and Physiology, School of Medicine and Dentistry, University of Rochester, 601 Elmwood Avenue, Rochester, NY 14642. E-mail:.