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Exposure of pancreatic islets of Langerhans to physiological concentrations of glucose leads to secretion of insulin in an oscillatory pattern. The oscillations in insulin secretion are associated with oscillations in cytosolic Ca2+ concentration ([Ca2+]c). Evidence suggests that the oscillations in [Ca2+]c and secretion are driven by oscillations in metabolism, but it is unclear whether metabolic oscillations are intrinsic to metabolism or require Ca2+ feedback. To address this question we explored the interaction of Ca2+ concentration and islet metabolism using simultaneous recordings of NAD(P)H autofluorescence and [Ca2+]c, in parallel with measurements of mitochondrial membrane potential (ΔΨm). All three parameters responded to 10 mm glucose with multiphasic dynamics culminating in slow oscillations with a period of ∼5 min. This was observed in ∼90% of islets examined from various mouse strains. NAD(P)H oscillations preceded those of [Ca2+]c, but their upstroke was often accelerated during the increase in [Ca2+]c, and Ca2+ influx was a prerequisite for their generation. Prolonged elevations of [Ca2+]c augmented NAD(P)H autofluorescence of islets in the presence of 3 mm glucose, but often lowered NAD(P)H autofluorescence of islets exposed to 10 mm glucose. Comparable rises in [Ca2+]c depolarized ΔΨm. The NAD(P)H lowering effect of an elevation of [Ca2+]c was reversed during inhibition of mitochondrial electron transport. These findings reveal the existence of slow oscillations in NAD(P)H autofluorescence in intact pancreatic islets, and suggest that they are shaped by Ca2+ concentration in a dynamic balance between activation of NADH-generating mitochondrial dehydrogenases and a Ca2+-induced decrease in NADH. We propose that a component of the latter reflects mitochondrial depolarization by Ca2+, which reduces respiratory control and consequently accelerates oxidation of NADH.
Glucose-induced insulin release from pancreatic islets of Langerhans depends on glucose uptake and metabolism in the β-cells, which elevates the cytosolic ATP/ADP ratio. This induces closure of ATP-sensitive potassium (KATP) channels in the plasma membrane (Misler et al. 1986; Malaisse & Sener, 1987), leading to cellular depolarization, voltage-gated calcium influx and, ultimately, the stimulation of calcium- and ATP-dependent insulin granule mobilization and exocytosis (Ashcroft & Rorsman, 1989; Misler et al. 1992b).
A hallmark of the normal insulin secretory response to glucose is its oscillatory nature, both in vivo (Lang et al. 1979; Porksen et al. 1995; Song et al. 2000) and in vitro in the perfused pancreas (Sturis et al. 1994) and isolated islets (Longo et al. 1991; Gilon et al. 1993; Bergsten et al. 1994; Westerlund & Bergsten, 2001). There is evidence that glucose-induced oscillations in islets and β-cells are promoted by periodic cycles in KATP channel activity (Larsson et al. 1996), indicating underlying fluctuations in ATP/ADP ratio and hence the involvement of dynamic metabolic events. Reports of metabolic oscillations in β-cells and islets are consistent with this (Longo et al. 1991; Jung et al. 2000; Ortsater et al. 2000; Ainscow & Rutter, 2002); however, the basis of such oscillations is debated. One possibility is that they are a consequence of enzymatic feedback mechanisms inherent to glycolysis (Tornheim, 1997; Juntti-Berggren et al. 2003). However, the picture may be more complex, as increases in cytosolic Ca2+ concentration ([Ca2+]c) evoked by glucose, or other agonists that elevate [Ca2+]c, are rapidly relayed to the mitochondria of β-cells in primary islets (Ainscow & Rutter, 2001) and insulin secreting cell lines (Rutter et al. 1993; Kennedy et al. 1996; Nakazaki et al. 1998). The resulting rise in the mitochondrial Ca2+ concentration ([Ca2+]m) may both augment oxidative phosphorylation, via activation of intramitochondrial dehydrogenases (McCormack et al. 1990a,b; Civelek et al. 1996b), and dissipate respiratory energy by the electrogenic uptake and futile cycling of Ca2+ across the inner mitochondrial membrane (Magnus & Keizer, 1998a; Nicholls & Ferguson, 2002). In the context of the present study, it is important to note that the intimate relationship between [Ca2+]c and [Ca2+]m in the β-cell also encompasses a close correlation of oscillatory Ca2+ dynamics in the two compartments (Nakazaki et al. 1998; Ainscow & Rutter, 2001).
In the present work, we demonstrate and characterize glucose-induced NAD(P)H and mitochondrial membrane potential oscillations in intact mouse islets, and establish a key role of Ca2+ in sustaining and shaping these oscillations. This is done, in part, by quantifying the temporal concordance of NAD(P)H and [Ca2+]c dynamics in islets exposed to glucose and agents that rapidly modulate Ca2+ influx. In particular, the simultaneous measurement of [Ca2+]c and NAD(P)H autofluorescence is achieved by using the Ca2+-sensitive dye Fura Red, whose fluorescent properties allow the joint detection of the relatively weak NAD(P)H autofluorescence signal.
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In the present study we examined the interrelation of glucose-induced [Ca2+]c, NAD(P)H autofluorescence and ΔΨm responses in intact mouse pancreatic islets. Using the long-wavelength Ca2+-sensitive fluorophore Fura Red, we were able to simultaneously monitor endogenous NAD(P)H autofluorescence and [Ca2+]c and, thus, to delineate their relationship during the distinct dynamic responses induced by glucose. Parallel measurements of ΔΨm were made, using average islet Rh123 fluorescence, to further elucidate the interaction of [Ca2+]c with islet respiration.
Our joint measurements show that the increase in islet NAD(P)H levels precedes the first [Ca2+]c rise above baseline by approximately 2 min and, hence, that the initiation of β-cell respiration and the subsequent elevation of the ATP/ADP ratio do not require, but rather lead to, the initial rise in [Ca2+]c. This temporal sequence of metabolic and ionic events has previously been observed in studies of glucose-stimulated islets and β-cells, in which [Ca2+]c and metabolic parameters have been recorded in parallel experiments (Pralong et al. 1990; Gilon & Henquin, 1992; Duchen et al. 1993; Detimary et al. 1998; Patterson et al. 2000) or simultaneously (Civelek et al. 1996a; Jung et al. 2000; Krippeit-Drews et al. 2000; Kindmark et al. 2001). The NAD(P)H response thus coincides with the onset of the characteristic phase 0 drop in [Ca2+]c, which is in agreement with the notion that phase 0 reflects the energy-dependent buffering of basal Ca2+ by intracellular organelles via thapsigargin-sensitive uptake (Gylfe, 1988; Chow et al. 1995).
Pralong et al. (1994) previously reported NAD(P)H fluctuations with a period of approximately 45 s in a subset (∼10%) of single isolated rat β-cells. With the exception of these recordings, periodic fluctuations in steady-state NAD(P)H levels have remained elusive in investigations of intact islets (Panten et al. 1973; Gilon & Henquin, 1992) and single β-cells (Krippeit-Drews et al. 2000; Kindmark et al. 2001). Contrary to the former islet studies, our findings demonstrate that intermediate concentrations of glucose reliably induce multiphasic NAD(P)H dynamics culminating in slow and regular oscillations. It remains unclear, however, exactly why we so consistently observed these islet oscillations while they have escaped detection in other studies. As islets from C57BL/6, Swiss-Webster and ob/ob mice showed similar behaviour, it is highly unlikely that the difference is directly related to the mouse strain under investigation.
Because of the elusive nature of oscillations of NAD(P)H autofluorescence, there has thus far been no direct information regarding the temporal concordance of oscillatory islet [Ca2+]c and NAD(P)H dynamics under physiological steady-state conditions. We established that at 10 mm glucose, the NAD(P)H oscillations have a period of roughly 5 min and are highly correlated with, but consistently lead, oscillations in [Ca2+]c, as demonstrated by their joint measurement. Furthermore we found that voltage-gated Ca2+ influx is a prerequisite for these slow NAD(P)H oscillations (Fig. 4). This is analogous to a dependence on extracellular Ca2+ observed for the faster NAD(P)H fluctuations of single rat β-cells (Pralong et al. 1994), an indication that fast and slow NAD(P)H dynamics to some degree share a common basis.
Similar to the situation in a variety of other cell types, mitochondria of β-cells sequester Ca2+ in response to increases in [Ca2+]c induced under physiological conditions (Rutter et al. 1993; Kennedy et al. 1996; Nakazaki et al. 1998; Ainscow & Rutter, 2001). This can affect the cellular energy state via stimulation of intramitochondrial Ca2+-sensitive dehydrogenases (McCormack et al. 1990a,b). Our recordings provide indirect evidence that, during physiological stimulation of pancreatic islets by glucose, [Ca2+]c dynamically activates these enzymes and increases the rate of TCA cycle flux. This manifests itself already during the initial phase of the response, where a transitory acceleration of the gradual NAD(P)H rise coincided with the peak in [Ca2+]c (Fig. 2). Previously it has been demonstrated that [Ca2+]c transients, induced by brief (30 s) pulses of a high concentration of KCl, initiate parallel pulses of NAD(P)H formation in quiescent rat β-cells in the presence of basal levels of glucose (Pralong et al. 1994). We found that extended periods of KCl-induced depolarization evokes similar metabolic stimulation of intact islets exposed to 3 mm glucose (Fig. 5A), which matches a rise in cytosolic ATP levels observed in rat islets depolarized under similar conditions (Ainscow & Rutter, 2001). While glucose-induced NAD(P)H oscillations could thus arise as [Ca2+]c oscillations are relayed to the mitochondria and periodically stimulate mCaDH, a number of our observations indicate that slow steady-state NAD(P)H oscillations are a manifestation of a more complex effect of elevated [Ca2+]c on the islet redox state. First, rises in NAD(P)H generally precede those of [Ca2+]c during spontaneous oscillations (Fig. 3). Second, while activation of mCaDH augments oscillations in NAD(P)H autofluorescence, prolonged rises in [Ca2+]c tend to decrease NAD(P)H levels (Fig. 5). Conceivably this effect comes into play during slow glucose-induced oscillations and contributes to the NAD(P)H autofluorescence downstroke. The lowering of NAD(P)H induced by Ca2+ is in contrast to the stimulation observed in presence of basal levels of glucose, and hinges on maintained flux through the electron transport chain (Fig. 6). Two principal mechanisms may account for this observation, namely the inhibition of β-cell phosphofructokinase, and hence glycolysis, by mitochondrially derived ATP (Yaney et al. 1995) and/or augmented NADH oxidation independent of changes in TCA cycle activity.
Hepatocytes stimulated by vasopressin show increases in NAD(P)H levels that return to prestimulatory levels despite sustained activation of pyruvate dehydrogenase (Robb-Gaspers et al. 1998). This is paralleled by an increase in the protonmotive force, and has therefore been attributed to NAD(P)H re-oxidation evoked by direct, Ca2+-dependent activation of the respiratory chain, possibly involving mitochondrial volume changes (Robb-Gaspers et al. 1998). In light of the present results we surmise that rises in islet NAD(P)H autofluorescence due to mCaDH activation are also countered by Ca2+-induced NADH oxidation. However, because islet ΔΨm is depolarized, rather than hyperpolarized, under conditions that lower NAD(P)H autofluorescence (cf. Figs 5 and 7A), the islet NAD(P)H re-oxidation is more likely to be a consequence of reduced respiratory control caused by the fall in the protonmotive force. The lowering of ΔΨm is presumably due to the continuous futile mitochondrial cycling of Ca2+ (Magnus & Keizer, 1998a; Krippeit-Drews et al. 2000; Kindmark et al. 2001). The NAD(P)H and ΔΨm measurements made here corroborate the presence of slow (> 3 min) oscillations in islet respiration, as previously documented by high resolution O2 recordings (Longo et al. 1991; Jung et al. 2000; Ortsater et al. 2000). In agreement with our findings, Jung et al. (2000) established that the O2 oscillations depended on Ca2+ influx. It is interesting that they also found that an oscillation in [Ca2+]c precedes that of O2 consumption. This is reconcilable with the observed oscillatory Ca2+–NAD(P)H phase relationship, as both a rise in NAD(P)H autofluorescence (mCaDH stimulation) and augmented NADH oxidation by the respiratory chain may be associated with increased respiratory flux and, consequently, a lowering of O2 levels.
Thus, our results suggest that [Ca2+]c plays an important role in the generation of slow oscillations in islet respiration. Nevertheless, it remains a possibility that Ca2+-independent oscillations exist that are merely too weak to be detected by the measurement techniques employed here. KATP channel activity in the β-cell has been reported to oscillate with periods between 2.5 and 4 min in the presence of substimulatory glucose levels where voltage-gated Ca2+ influx is not activated (Dryselius et al. 1994). There is conflicting evidence regarding the role of Ca2+ in sustaining O2 oscillations. In contrast to the findings of Jung et al. (2000), other islet studies (Longo et al. 1991; Ortsater et al. 2000) and measurements from insulin-secreting clonal (HIT) β-cells (Porterfield et al. 2000) have identified low-amplitude O2 oscillations in the presence of basal levels of glucose and under Ca2+-free conditions. The latter were, however, significantly amplified upon elevation of the Ca2+ concentration.
The phase relationship observed in our steady-state recordings is consistent with a scenario where metabolic oscillations, via regulation of the KATP channel, induce slow oscillations of [Ca2+]c and, ultimately, insulin secretion. Metabolic oscillations in the β-cell have been proposed to arise from intrinsic glycolytic mechanisms involving the allosteric feedback-activation of phosphofructokinase (Tornheim, 1997; Juntti-Berggren et al. 2003). That glycolytic flux is oscillatory has been demonstrated by recordings of oscillations in islet glucose consumption (Jung et al. 2000) and islet lactate release (Chou et al. 1992). However, a purely glycolytic basis of these oscillations is in apparent conflict with the Ca2+ requirement for oscillations in NAD(P)H and ΔΨm, which rather supports the presence of a full feedback loop where oscillations in [Ca2+]c and metabolism arise through a mutual interdependence.
Several mechanisms have been proposed by which [Ca2+]c feedback may reduce cellular ATP levels and re-activate the KATP channels (reviewed by Kennedy et al. 2002). These include: (i) Ca2+-stimulated ATP hydrolysis (Detimary et al. 1998; Ainscow & Rutter, 2002); (ii) reduced ATP synthesis due to futile mitochondrial Ca2+ cycling (Magnus & Keizer, 1998a; Krippeit-Drews et al. 2000; Kindmark et al. 2001); and (iii) Ca2+-dependent feedback inhibition of glycolytic flux (Jung et al. 2000). Our results are most directly compatible with the second of these possibilities, but do not rule out the other ATP-lowering mechanisms. When considering this, it should also be kept in mind that both futile mitochondrial cycling of Ca2+ and increased cytosolic ATP consumption may in fact accelerate respiration by attenuating respiratory control. Consequently, for the Ca2+-dependent negative feedback mechanisms to repolarize the β-cell, it is necessary that the sum of their effects exceed this compensatory increase in oxidative phosphorylation.
Conversely, the putative impact of the recorded NAD(P)H autofluorescence and ΔΨm oscillations on islet [Ca2+]c and secretion dynamics depends on their ability to effectively change the rate of oxidative phosphorylation. Based on detailed mathematical modelling of β-cell mitochondrial metabolism and calcium handling it has been predicted that ΔΨm oscillations with an amplitude of less than 1 mV provide adequate regulation of ATP synthesis to induce KATP channel-dependent bursting electrical behaviour (Magnus & Keizer, 1998a,b). In the model, the size of these ΔΨm fluctuations is approximately 10% of the total mitochondrial hyperpolarization observed upon a simulated glucose rise, and the corresponding mitochondrial NADH oscillations have an amplitude of roughly 20% of the total glucose-induced NADH rise (Magnus & Keizer, 1998b). These theoretical estimates are comparable to the experimental observations in the present study. This supports the possibility that the recorded metabolic oscillations are of functional consequence and, hence, that the interactions between Ca2+ concentration and respiration are in fact of a reciprocal nature. A more quantitative verification of these model predictions is an important task to be addressed in future experiments.
In addition to the scenarios discussed above, there is a composite model in which an intrinsic slow glycolytic oscillator co-exists with indirect Ca2+-dependent modulation of glycolytic flux. This alternative model has been constructed and examined mathematically (Bertram et al. 2004) and extending this work promises to provide further insights into the fundamental, yet remarkably complex, questions of how Ca2+, glycolysis and mitochondria interact to regulate β-cell oscillatory behaviour.
In summary, the present work has established that glucose consistently induces slow NAD(P)H autofluorescence oscillations in intact mouse pancreatic islets, and clarified how these arise through interaction with changes in Ca2+ concentration. The data indicate that [Ca2+]c shapes the NAD(P)H oscillations by first accelerating the reduction of pyridine nucleotides and then invoking the subsequent drop in NAD(P)H autofluorescence, in part by stimulating NADH oxidation in the electron transport chain. When [Ca2+]c falls at the end of an oscillation, NAD(P)H levels can once again increase to complete the periodic cycle. Moreover, our findings provide evidence to support the hypothesis that slow glucose-induced oscillations in islet respiration depend on feedback interactions between ionic and metabolic events in the pancreatic β-cell. These dynamic cross-talk mechanisms are therefore likely to be essential for normal pulsatile insulin release dynamics and may potentially be compromised when this pulsatility of insulin release is impaired in the development of type 2 diabetes.